Encyclopedia of Geobiology

ENCYCLOPEDIA of GEOBIOLOGY

Encyclopedia of Earth Sciences Series ENCYCLOPEDIA OF GEOBIOLOGY Volume Editors

Joachim Reitner is Professor of Paleontology, Head of the Department of Geobiology, and Managing Director of the Museum, Collections and Geopark, at the University of Göttingen, Germany. He is also Editor-in-Chief of Lecture Notes in Earth Sciences (Springer), Co-Editor of Facies (Springer), and Associate Editor of the Geomicrobiology Journal (Taylor & Francis). Dr. Reitner’s research focuses on the interplay between organisms and their metabolic processes with various abiotic parameters. Many geological processes can be understood as geo-physiological processes, allowing chemical reactions to proceed that would never occur under standard thermodynamic conditions. Therefore, a major thrust of Dr. Reitner’s research is the investigation of the evolution of these processes, which are visible in biosignatures and biomineralization patterns, and in their interaction with biogeochemical cycles. Among his many honors and accolades, Dr. Reitner is the recipient of the G. W. Leibniz Award from the Deutsche Forschungsgemeinschaft. Volker Thiel is Professor of Organic Geochemistry in the Geoscience Center at the University of Göttingen, Germany. Dr. Thiel has been involved in geobiological research for some 15 years, with a focus on the use of organic molecules as chemical tracers (biomarkers) for biogeochemical pathways. His research interests include lipid biomarkers as indicators for biogeochemical processes, molecular fossils, biological formation, and turnover of methane, and microbial control on mineral formation. The results of his studies have significantly contributed to the characterization of microbial processes associated with methane turnover in modern and ancient environments. Much of Dr. Thiel’s current work is devoted to new approaches to enhance the spatial resolution of biomarker analysis in geobiological systems. He is member of the Editorial Board of the journal Geobiology (Wiley-Blackwell).

Editorial Board

Hans-Joachim Fritz Courant Research Centre Geobiology University of Göttingen Goldschmidtstr.3 37077 Göttingen Germany

Pamela Reid Rosenstiel School of Marine and Atmospheric Science University of Miami 4600 Rickenbacker Cswy Miami FL 33149 USA

Andreas Kappler Center for Applied Geoscience (ZAG) Sigwartstraße 10 72076 Tübingen Germany

Xingliang Zhang Department of Geology Northwest University Xian 710069 China

Kurt O. Konhauser Department of Earth and Atmospheric Sciences University of Alberta, Edmonton Alberta, T6G 2E3 Canada

Aims of the Series

The Encyclopedia of Earth Sciences Series provides comprehensive and authoritative coverage of all the main areas in the Earth Sciences. Each volume comprises a focused and carefully chosen collection of contributions from leading names in the subject, with copious illustrations and reference lists. These books represent one of the world’s leading resources for the Earth Sciences community. Previous volumes are being updated and new works published so that the volumes will continue to be essential reading for all professional earth scientists, geologists, geophysicists, climatologists, and oceanographers as well as for teachers and students. See http://www.springer.com for a current list of titles in the Encyclopedia of Earth Sciences Series. Go to http://www.springerlink.com/reference-works/ to visit the “Encyclopedia of Earth Sciences Series” on-line.

About the Series Editor

Professor Charles W. Finkl has edited and/or contributed to more than 8 volumes in the Encyclopedia of Earth Sciences Series. For the past 25 years he has been the Executive Director of the Coastal Education & Research Foundation (CERF) and Editor-in-Chief of the international Journal of Coastal Research. In addition to these duties, he is Research Professor at Florida Atlantic University in Boca Raton, Florida, USA. He is a graduate of the University of Western Australia (Perth) and previously worked for a wholly owned Australian subsidiary of the International Nickel Company of Canada (INCO). During his career, he acquired field experience in Australia; the Caribbean; South America; SW Pacific islands; southern Africa; Western Europe; and the Pacific Northwest, Midwest, and Southeast USA.

Founding Series Editor

Professor Rhodes W. Fairbridge (deceased) has edited more than 24 Encyclopedias in the Earth Sciences Series. During his career he has worked as a petroleum geologist in the Middle East, been a WW II intelligence officer in the SW Pacific and led expeditions to the Sahara, Arctic Canada, Arctic Scandinavia, Brazil and New Guinea. He was Emeritus Professor of Geology at Columbia University and was affiliated with the Goddard Institute for Space Studies.

ENCYCLOPEDIA OF EARTH SCIENCES SERIES

ENCYCLOPEDIA of GEOBIOLOGY edited by

JOACHIM REITNER VOLKER THIEL University of Göttingen Germany

Library of Congress Control Number: 2010936497

ISBN: 978-1-4020-9211-4 This publication is available also as: Electronic publication under ISBN 978-1-4020-9212-1 and Print and electronic bundle under ISBN 978-1-4020-9213-8

Published by Springer P.O. Box 17, 3300 AA Dordrecht, The Netherlands

Printed on acid-free paper

Cover illustration: Iron- and sulfide-oxidizing microbial mats in the Äspö Hard Rock Laboratory, Oskarshamn, Sweden (photograph by Joachim Reitner) Every effort has been made to contact the copyright holders of the figures and tables which have been reproduced from other sources. Anyone who has not been properly credited is requested to contact the publishers, so that due acknowledgment may be made in subsequent editions.

All Rights Reserved © Springer Science þ Business Media B.V. 2011 No part of this work may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording, or otherwise, without written permission from the Publisher, with the exception of any material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work.

Contents

Contributors Preface

xiii xxvii

Animal Biocalcification, Evolution Gert Wörheide and Daniel J. Jackson

53

Acetogens Kirsten Küsel and Harold L. Drake

1

Animal Skeletons, Advent Guoxiang Li, Maoyan Zhu and Zhe Chen

58

Acid Rock Drainage Lesley A. Warren

5

Archaea Volker Thiel

64

Acidophiles

8

Arsenic John F. Stolz and Ronald S. Oremland

69

Acritarchs

8

69

Aerobic Metabolism Heribert Cypionka

8

Asteroid and Comet Impacts Charles S. Cockell Astrobiology Jack D. Farmer

73

Bacteria Michael Hoppert

81

Bacterioplankton Thomas Pommier

89 92

Algae (Eukaryotic) Thomas Friedl, Nicole Brinkmann and Kathrin I. Mohr

10

Alkalinity Andreas Reimer and Gernot Arp

20

Amber Eugenio Ragazzi and Alexander R. Schmidt

24

Anaerobic Oxidation of Methane with Sulfate Katrin Knittel and Antje Boetius

36

Banded Iron Formations Nicole R. Posth, Kurt O. Konhauser and Andreas Kappler

Anaerobic Transformation Processes, Microbiology Bernhard Schink

48

Basalt (Glass, Endoliths) Ingunn H. Thorseth

103

Anammox

53

Beggiatoa Heide N. Schulz-Vogt

111

Entries without author names are glossary terms

vi

CONTENTS

Biodeterioration (of Stone) Christina Beimforde

112

Calcite Precipitation, Microbially Induced Tanja Bosak

223

Bioerosion Aline Tribollet, Gudrun Radtke and Stjepko Golubic

117

Calcium Biogeochemistry Anton Eisenhauer

227

Biofilms Joachim Reitner

134

Cap Carbonates Joachim Reitner

229

Biofilms and Fossilization Joachim Reitner

136

Carbon (Organic, Cycling) Amber K. Hardison and Elizabeth A. Canuel

230

Biogeochemical Cycles

137

Carbon (Organic, Degradation) Steven T. Petsch

234

Carbon Cycle

238

137

Carbon Isotopes

238

Biological Volcanic Rock Weathering Charles S. Cockell

143

Carbonate Environments Eberhard Gischler

238

Biomarkers (Molecular Fossils) Jochen J. Brocks and Kliti Grice

147

Carbonates Martin Dietzel

261

Cathodoluminescence Microscopy Walter Vortisch

266

Chemolithotrophy Volker Thiel

271

Biological Control on Diagenesis: Influence of Bacteria and Relevance to Ocean Acidification Fred T. Mackenzie and Andreas J. Andersson

Biomarkers (Organic, Compound-Specific Isotopes) Kliti Grice and Jochen J. Brocks

167

Biomining (Mineral Bioleaching, Mineral Biooxidation) Douglas Eric Rawlings

182

Cherts Volker Thiel

272

Bioprotection Carlos Rodriguez-Navarro, Maria T. González-Muñoz, Concepción Jimenez-Lopez and Manuel Rodriguez-Gallego

185

Chondrites

273

Chroococcidiopsis Burkhard Büdel

273

Biosignatures in Rocks Frances Westall and Barbara Cavalazzi

189

Clay Authigenesis, Bacterial Kurt O. Konhauser

274

Biosilicification

201

277

Black Shales Wolfgang Oschmann

201

Coccolithophores Volker Thiel

278

Breakup of Rodinia Zheng-Xiang Li

206

Cold Seeps Robert G. Jenkins Comets

290

Calcareous Algae

211

Commensalism

290

Calcification

211

Community

290

Calcified Cyanobacteria Robert Riding

211

Copper Stephan M. Kraemer

290

CONTENTS

vii

Cosmic Molecular Clouds Joachim Reitner

292

Endoliths Bettina Weber and Burkhard Büdel

348

Critical Intervals in Earth History Frank Wiese and Joachim Reitner

293

Endosymbiosis

355

Cryobiosphere

306

Evaporites

355

Cyanobacteria Kathrin I. Mohr, Nicole Brinkmann and Thomas Friedl

306

Exoenzymes Kathrin Riedel and Alexander Grunau

355 359

Deep Biosphere of Salt Deposits Helga Stan-Lotter and Sergiu Fendrihan

313

Extracellular Polymeric Substances (EPS) Alan W. Decho

362

Deep Biosphere of Sediments

317

Extreme Environments Volker Thiel

Deep Biosphere of the Oceanic Deep Sea Kristina Rathsack, Nadia-Valérie Quéric and Joachim Reitner

317

Fe(II)-Oxidizing Prokaryotes Kristina L. Straub

367 370

Deep Fluids

322

Fe(III)-Reducing Prokaryotes Kristina L. Straub

Degradation (of Organic Matter)

322

Fermentation

373

Denitrification

322

373

Desert Varnish Randall S. Perry

322

Fluorescence In Situ Hybridization (FISH) Natuschka M. Lee, Daniela B. Meisinger, Michael Schmid, Michael Rothballer and Frank E. Löffler

Detachment

325

Foraminifera Alexander V. Altenbach

393

Diatoms Nicole Brinkmann, Thomas Friedl and Kathrin I. Mohr

326

396

Dinoflagellates

331

Frutexites Marta Rodríguez-Martínez, Christine Heim, Nadia-Valérie Quéric and Joachim Reitner

Divalent Earth Alkaline Cations in Seawater Anton Eisenhauer

331

Fungi and Lichens Bettina Weber and Burkhard Büdel

401

Diversity

336

Gallionella Karsten Pedersen

411

Dolomite, Microbial Jennifer A. Roberts and Paul A. Kenward

336

412

Early Earth

341

Geobacter Kristina L. Straub

Early Precambrian Eukaryotes Joachim Reitner

341

Geochronology Volker Liebetrau

413

Ecological Niche

342

Geomycology Geoffrey M. Gadd

416

Ediacaran Biota Dmitriy Grazhdankin

342

Geyserite

433

Entries without author names are glossary terms

viii

CONTENTS

Glass

433

Glaucophytes

433

Gold Veit-Enno Hoffmann

433

Gondwanaland, Formation Joseph G. Meert

434

Great Oxygenation Event (GOE)

436

Green Algae

436

Guild

436

Habitat

437

Halobacteria – Halophiles Helga Stan-Lotter

437

Haptophytes

441

Heavy Metals

441

Histology Michael Gudo, Gerta Fleissner and Guenther Fleissner

441

Hot Springs and Geysers Brian Jones and Robin W. Renaut

Isotope Fractionation (Metal) Ariel D. Anbar and Silke Severmann

502

Isotopes (Methods)

511

Isotopes and Geobiology Jochen Hoefs

511

Isotopes, Radiogenic Bent T. Hansen

516

Karst Ecosystems Annette S. Engel

521

Lateral Gene Transfer Hans-Joachim Fritz

533

Leptothrix David Emerson

535

Magnetotactic Bacteria Mihály Pósfai

537

Manganese (Sedimentary Carbonates and Sulfides) 541 Michael E. Böttcher Mass Extinctions, Phanerozoic Joachim Reitner

543

447

Mat-Related Sedimentary Structures Hubertus Porada

547

Hydrogen Tori M. Hoehler

451

Metagenomics Wolfgang Liebl

553

Hydrothermal Environments, Fossil Joachim Reitner

454

Metalloenzymes Michael Hoppert

558

Hydrothermal Environments, Marine Gilberto E. Flores and Anna-Louise Reysenbach

456

Metallogenium Joachim Reitner

563

Hydrothermal Environments, Terrestrial Robin W. Renaut and Brian Jones

467

Metals, Acquisition by Marine Bacteria Alison Butler and Vanessa V. Homann

565

Hypersaline Environments

479

Meteoritics Mark A. Sephton

568

Ichnology Murray Gingras and Kurt O. Konhauser

481

Methane Oxidation (Aerobic) Helmut Bürgmann

575

Immunolocalization Michael Hoppert and Christoph Wrede

482

Methane, Origin Carsten J. Schubert

578

Iron Isotopes

486

Methanogens

586

Iron Sulfide Formation Jürgen Schieber

486

Microbial Biomineralization Christine Heim

586

CONTENTS

ix

Microbial Communities, Structure, and Function Michael W. Friedrich

592

Nickel, Biology Martin Krüger

684

Microbial Degradation Erika Kothe

596

Nitrification

685

Microbial Ecology of Submarine Caves Francesco Canganella and Giovanna Bianconi

599

Nitrogen Volker Thiel

686

Microbial Mats Joachim Reitner

606

Nitrogen Fixation

690 691

Microbial Silicification – Bacteria (or Passive) Kurt O. Konhauser and Brian Jones

608

Ores, Microbial Precipitation and Oxidation Beda A. Hofmann Organic Carbon

697

Microbial Surface Reactivity David A. Fowle and Kurt O. Konhauser

614

Organomineralization Christian Défarge

697

Microbialites, Modern Christophe Dupraz, R. Pamela Reid and Pieter T. Visscher

617

Origin of Life Michael J. Russell

701

Microbialites, Stromatolites, and Thrombolites Robert Riding

635

Origins of the Metazoa Daniel J. Jackson

716

Microbial-Metal Binding Kurt O. Konhauser and David A. Fowle

654

Parasitism

721 721

Microbiocorrosion Aline Tribollet, Stjepko Golubic, Gudrun Radtke and Joachim Reitner

657

Pedogenic Carbonates Eric P. Verrecchia Permafrost Microbiology David A. Gilichinsky and Elizaveta M. Rivkina

726

658

Phosphorus, Phosphorites Karl B. Föllmi

732

Molar-tooth Structure Brian R. Pratt

662

Photosynthesis Kerstin Schmidt

736

Moonmilk Joachim Reitner

666

Piezophilic Bacteria Jiasong Fang and Li Zhang

738

Mud Mounds Marta Rodríguez-Martínez

667

Pore Waters Sabine Kasten

742

Mutualism

675

Protozoa (Heterotroph, Eukaryotic) Jens Boenigk

746

Mycorrhizae

675

Pyrite Oxidation

750

Nan(n)obacteria Muriel Pacton and Georges E. Gorin

677

Radioactivity (Natural) Beda A. Hofmann

751

Nanocrystals, Microbially Induced Susan Glasauer

681

Radiolaria Volker Thiel

754

Microsensors for Sediments, Microbial Mats, and Biofilms Dirk de Beer

Entries without author names are glossary terms

x

CONTENTS

Soda Lakes Stephan Kempe and Jozef Kazmierczak

824

761

Soda Ocean Hypothesis Stephan Kempe and Jozef Kazmierczak

829

Reefs

762

Soils Erika Kothe

833

Remineralization (of Organic Matter)

763

Species (Microbial)

836

Rhodophyta

763

836

RNA-World

763

Speleothems Roman Aubrecht

Saline Lakes Carol D. Litchfield

765

Sponges (Porifera) and Sponge Microbes Friederike Hoffmann and Marie-Lise Schläppy

840

Salinity History of the Earth’s Ocean L. Paul Knauth

769

Stromatactis Roman Aubrecht

847

Stromatolites

850

772

Subsurface Filamentous Fabrics Beda A. Hofmann

851

Sediment Diagenesis – Biologically Controlled Kurt O. Konhauser, Murray K. Gingras and Andreas Kappler

777

Sulfate-Reducing Bacteria Heribert Cypionka

853

Selenium John F. Stolz and Ronald S. Oremland

784

Sulfide Mineral Oxidation D. Kirk Nordstrom

856

Shales Jürgen Schieber

785

Sulfur Cycle Michael E. Böttcher

859

Shewanella Nadia-Valérie Quéric

791

Sulfur Isotopes Michael E. Böttcher

864

Siderite Volker Thiel

792

Symbiosis Sharmishtha Dattagupta and Frank Zielinski

866

Siderophores Stephan M. Kraemer

793

Syntrophy

870 871

Silica Biomineralization, Sponges Hermann Ehrlich

796

Terrestrial Deep Biosphere Christine Heim

876

Silicoflagellates

808

Thioester World Joachim Reitner

Sinter Robin W. Renaut and Brian Jones

808

Thiomargarita Heide N. Schulz-Vogt

877

Skeleton

814

Thiotrophic Bacteria Heide N. Schulz-Vogt

877

Small Shelly Fossils

814

Thrombolites

880

Snowball Earth Paul F. Hoffman

814

Tidal Flats Meinhard Simon

880

Raman Microscopy (Confocal) Jan Toporski, Thomas Dieing and Christine Heim

754

Reduction Spheroids Beda A. Hofmann

Scanning Probe Microscopy (Includes Atomic Force Microscopy) Michael Hoppert

CONTENTS

xi

TOF-SIMS Peter Sjövall and Jukka Lausmaa

883

Whale and Wood Falls Steffen Kiel

901

Trace Fossils: Neoproterozoic Sören Jensen, James G. Gehling and Mary L. Droser

886

Zinc Matthias Labrenz and Gregory K. Druschel

905

Tufa, Freshwater Akihiro Kano

889

Author Index

909

Subject Index

911

Waulsortian Mud Mounds Marta Rodríguez-Martínez

893

Entries without author names are glossary terms

Contributors

Alexander V. Altenbach Ludwig-Maximilians-University Munich Richard-Wagner-Str. 10 80333 Munich Germany [email protected] Ariel D. Anbar School of Earth and Space Exploration and Department of Chemistry and Biochemistry Arizona State University Tempe, AZ USA [email protected] Andreas J. Andersson Bermuda Institute of Ocean Sciences 17 Biological Station Ferry Reach, St. George’s Bermuda Gernot Arp Geobiology Group Geoscience Center University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Roman Aubrecht Department of Geology and Paleontology Faculty of Natural Sciences Comenius University Mlynská dolina-G 842 15 Bratislava Slovakia [email protected]

Christina Beimforde Courant Research Centre Geobiology University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Giovanna Bianconi Department of Agrobiology and Agrochemistry University of Tuscia Viterbo Italy [email protected] Jens Boenigk Institute for Limnology Austrian Academy of Sciences Mondseestr. 9 5310 Mondsee Austria [email protected] Antje Boetius Department of Molecular Ecology Max Planck Institute for Marine Microbiology Celsiusstrasse 1 28359 Bremen Germany [email protected] Tanja Bosak Department of Earth Atmospheric and Planetary Sciences Massachusetts Institute of Technology Cambridge, MA 02139 USA [email protected]

xiv

CONTRIBUTORS

Michael E. Böttcher Geochemistry & Stable Isotope Geochemistry Marine Geology Section Leibniz Institute for Baltic Sea Research 18119 Warnemünde Germany [email protected]

Elizabeth A. Canuel College of William and Mary Virginia Institute of Marine Science P.O. Box 1346 Gloucester Point, VA 23062 USA [email protected]

Nicole Brinkmann Geobiology Group Geoscience Center University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected]

Barbara Cavalazzi Centre de Biophysique Moléculaire CNRS Université de Orléans Rue Charles Sadron 45071 Orléans France

Jochen J. Brocks Research School of Earth Sciences The Australian National University Building 61, Mills Road Canberra, ACT 0200 Australia [email protected] Burkhard Büdel Plant Ecology and Systematics Department of Biology University of Kaiserslautern P.O. Box 3049 67653 Kaiserslautern Germany [email protected] Helmut Bürgmann Department of Surface Waters Eawag, Swiss Federal Institute of Aquatic Science and Technology Seestrasse 79 6047 Kastanienbaum Switzerland [email protected] Alison Butler Department of Chemistry University of California Santa Barbara, CA 93106-9510 USA [email protected] Francesco Canganella Department of Agrobiology and Agrochemistry University of Tuscia Viterbo Italy [email protected]

Zhe Chen State Key Laboratory of Palaeobiology and Stratigraphy Chinese Academy of Sciences 210008 Nanjing China Charles S. Cockell Planetary and Space Sciences Research Institute Open University Milton Keynes, MK7 6AA UK [email protected] Heribert Cypionka University of Oldenburg 26111 Oldenburg Germany [email protected] Sharmishtha Dattagupta Courant Research Centre Geobiology University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Dirk de Beer Microsensor Research Group Max Planck Institute for Marine Microbiology Celsiusstrasse 1 28359 Bremen Germany [email protected] Alan W. Decho Department of Environmental Health Sciences Arnold School of Public Health University of South Carolina Columbia, SC 29208 USA [email protected]

CONTRIBUTORS

Christian Défarge Institut des Sciences de la Terre d’Orléans Unité Mixte de Recherche 6113 Université d’Orléans and Université François-Rabelais de Tours Ecole Polytechnique de l’Université d’Orléans 8 rue Léonard de Vinci 45072 Orléans France [email protected] Thomas Dieing Wissenschaftliche Instrumente und Technologie GmbH Lise-Meitner-Strasse 6 89081 Ulm Germany [email protected]

Hermann Ehrlich Biominerals, Biocomposites & Biomimetics Group Institute of Bioanalytical Chemistry Dresden University of Technology Bergstr. 66 1062 Dresden Germany [email protected] Anton Eisenhauer IFM-GEOMAR Leibniz-Institut für Meereswissenschaften Christian Albrechts Universität zu Kiel Wischhofstr. 1-3 24148 Kiel Germany [email protected]

Martin Dietzel Institute of Applied Geosciences Graz University of Technology 8010 Graz Austria [email protected]

David Emerson Bigelow Laboratory for Ocean Sciences West Boothbay Harbor, ME 04575 USA [email protected]

Harold L. Drake Department of Ecological Microbiology University of Bayreuth 95440 Bayreuth Germany [email protected]

Annette S. Engel Department of Geology and Geophysics Louisiana State University E235 Howe-Russell Geoscience Complex Baton Rouge, LA 70803 USA [email protected]

Mary L. Droser Department of Earth Sciences University of California Riverside, CA 92521 USA [email protected] Gregory K. Druschel Department of Geology University of Vermont 321 Delehanty Hall 180 Colchester Ave. Burlington, VT 05405 USA [email protected] Christophe Dupraz Center for Integrative Geosciences – Marine Sciences University of Connecticut 354 Mansfield Road Storrs, CT 06269-2045 USA [email protected]

xv

Jiasong Fang College of Natural and Computational Sciences Hawaii Pacific University Kaneohe, HI 96744 USA [email protected] Jack D. Farmer School of Earth and Space Exploration Arizona State University Tempe, AZ USA [email protected] Sergiu Fendrihan Bioresource Center and Advanced Research Association Aleea Istru nr. 2C, bloc A14B sc. 8, et. 2, apt. 113, sect. 6 061912 Bucharest Romania [email protected]; [email protected]

xvi

CONTRIBUTORS

Gerta Fleissner Institute for Cell Biology and Neurosciences Goethe-University Frankfurt Siesmayerstr. 70 60323 Frankfurt a. M. Germany [email protected]

Hans-Joachim Fritz Courant Research Centre Geobiology University of Göttingen Goldschmidstr. 3 37077 Göttingen Germany [email protected]

Guenther Fleissner Institute for Cell Biology and Neurosciences Goethe-University Frankfurt Siesmayerstr. 70 60323 Frankfurt a. M. Germany [email protected]

Geoffrey M. Gadd Division of Molecular Microbiology College of Life Sciences University of Dundee Dundee DD1 5EH Scotland UK [email protected]

Gilberto E. Flores Department of Biology Portland State University 1719 SW 10th Avenue Portland, OR 97207 USA [email protected]

James G. Gehling South Australian Museum Adelaide, SA 5000 Australia [email protected]

Karl B. Föllmi Institute of Geology and Paleontology University of Lausanne 2015 Lausanne Switzerland [email protected]

David A. Gilichinsky Institute of Physicochemical & Biological Problems in Soil Science Russian Academy of Sciences Pushchino Russia [email protected]

David A. Fowle Department of Earth and Atmospheric Sciences University of Alberta Edmonton, Alberta T6G 2E3 Canada [email protected]

Murray K. Gingras Department of Earth and Atmospheric Sciences University of Alberta Edmonton, Alberta T6G 2E3 Canada [email protected]

Thomas Friedl Albrecht-von-Haller-Institute for Plant Sciences University of Göttingen Untere Karspüle 2 37073 Göttingen Germany [email protected]

Eberhard Gischler Institut für Geowissenschaften Facheinheit Paläontologie J.W. Goethe-Universität Altenhöferallee 1 60438 Frankfurt am Main Germany [email protected]

Michael W. Friedrich Microbial Ecophysiology Faculty of Biology/Chemistry University of Bremen 28359 Bremen Germany [email protected]

Susan Glasauer Department of Land Resource Science University of Guelph Guelph, Ontario N1G 2W1 Canada [email protected]

CONTRIBUTORS

xvii

Stjepko Golubic Department of Biology Boston University 5 Cummington Street Boston, MA 02215 USA [email protected]

Michael Gudo Morphisto Evolutionsforschung und Anwendung GmbH Institut für Evolutionswissenschaften Weismüllerstr. 45 60314 Frankfurt am Main Germany [email protected]

Maria T. González-Muñoz Departamento de Mineralogía y Petrología Universidad de Granada Fuentenueva s/n 18002 Granada Spain [email protected]

Bent T. Hansen Department of Isotope Geology University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected]

Georges E. Gorin Department of Geology-Paleontology University of Geneva 13 rue des Maraîchers 1205 Geneva Switzerland [email protected]

Amber K. Hardison College of William and Mary Virginia Institute of Marine Science P.O. Box 1346 Gloucester Point, VA 23062 USA [email protected]

Dmitriy Grazhdankin Division of Precambrian and Cambrian Paleontology and Stratigraphy Russian Academy of Sciences Prospekt Akademika Koptyga 3 Novosibirsk 630090 Russia [email protected] Kliti Grice WA Organic and Isotope Geochemistry Centre Department of Applied Chemistry Curtin University of Technology Perth, WA 6845 Australia [email protected] Alexander Grunau Department of Microbiology Institute of Plant Biology University of Zurich Winterthurerstrasse 190 8057 Zurich Switzerland [email protected]

Christine Heim Geobiology Group Geoscience Centre University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Jochen Hoefs Department of Isotope Geology University of Göttingen Goldschmidtstr. 1 37077 Göttingen Germany [email protected] Tori M. Hoehler Exobiology Branch NASA-Ames Research Center Mail Stop 239-4 Moffett Field, CA 94035 USA [email protected]

xviii

CONTRIBUTORS

Paul F. Hoffman Department of Earth & Planetary Sciences Harvard University Cambridge, MA USA [email protected] Friederike Hoffmann Sars International Centre for Marine Molecular Biology 5008 Bergen Norway [email protected] Veit-Enno Hoffmann Department of Sedimentology and Environmental Geology University of Göttingen 37077 Göttingen Germany [email protected] Beda A. Hofmann Earth Science Department Natural History Museum Bern Bernastrasse 15 3005 Bern Switzerland [email protected] Vanessa V. Homann Department of Chemistry University of California Santa Barbara, CA 93106-9510 USA [email protected] Michael Hoppert Institut für Mikrobiologie und Genetik University of Göttingen Grisebachstraße 8 37077 Göttingen Germany [email protected] Daniel J. Jackson Courant Research Centre Geobiology University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected]

Robert G. Jenkins Faculty of Education and Human Sciences Yokohama National University 79-1 Tokiwadai Hodogaya-ku Yokohama City 240-8501 Japan [email protected] Sören Jensen Area de Paleontología Universidad de Extremadura 06006 Badajoz Spain [email protected] Concepción Jimenez-Lopez Departamento de Mineralogía y Petrología Universidad de Granada Fuentenueva s/n 18002 Granada Spain [email protected] Brian Jones Department of Earth and Atmospheric Sciences University of Alberta 1-26 Earth Sciences Building Edmonton, Alberta T6G 2E3 Canada [email protected] Akihiro Kano Department of Evolution of the Earth and the Environment Graduate School of Social and Cultural Studies Kyushu University Fukuoka 819-0395 Japan [email protected] Andreas Kappler Geomicrobiology Group Center for Applied Geoscience University of Tübingen Sigwartstrasse 10 72072 Tübingen Germany [email protected] Sabine Kasten Alfred Wegener Institute for Polar and Marine Research Am Handelshafen 12 27570 Bremerhaven Germany [email protected]

CONTRIBUTORS

Jozef Kazmierczak Institute of Paleobiology Polish Academy of Sciences ul. Twarda 51/55 00-818 Warszawa Poland [email protected]

Erika Kothe Institute of Microbiology Friedrich Schiller University Jena Neugasse 25 07743 Jena Germany [email protected]

Stephan Kempe Department of Physical Geology and Global Cycles Institute for Applied Geosciences University of Technology Schnittspahnstr. 9 64287 Darmstadt Germany [email protected]

Stephan M. Kraemer Department of Environmental Geosciences University of Vienna 1090 Vienna Austria [email protected]

Paul A. Kenward Department of Geology University of Kansas Lawrence, KS 66045-7613 USA [email protected] Steffen Kiel Geoscience Center Geobiology Group University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] L. Paul Knauth Arizona State University Box 871404 Tempe, AZ 85287-1404 USA [email protected] Katrin Knittel Department of Molecular Ecology Max Planck Institute for Marine Microbiology Celsiusstrasse 1 28359 Bremen Germany [email protected] Kurt O. Konhauser Department of Earth and Atmospheric Sciences University of Alberta Edmonton, Alberta T6G 2E3 Canada [email protected]

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Martin Krüger Geomicrobiology Federal Institute for Geosciences and Natural Resources Stilleweg 2 30655 Hannover Germany [email protected] Kirsten Küsel Institute of Ecology Friedrich Schiller University Jena 07743 Jena Germany [email protected] Matthias Labrenz IOW-Leibniz Institute for Baltic Sea Research Section Biology Seestrasse 15 18119 Rostock-Warnemuende Germany [email protected] Jukka Lausmaa SP Technical Research Institute of Sweden Chemistry and Materials Technology Box 857, SE-501 15 Borås Sweden [email protected] Natuschka M. Lee Department of Microbiology Technical University of Munich Emil-Ramann-Str. 4 85354 Freising-Weihenstephan Germany [email protected]

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CONTRIBUTORS

Guoxiang Li State Key Laboratory of Palaeobiology and Stratigraphy Chinese Academy of Sciences 210008 Nanjing China [email protected]

Joseph G. Meert Geological Sciences University of Florida 274 Williamson Hall Gainesville, FL USA [email protected]; [email protected]

Zheng-Xiang Li Department of Applied Geology Curtin University of Technology GPO Box U1987 Perth, WA 6845 Australia [email protected]

Daniela B. Meisinger Department of Microbiology Technical University of Munich 85354 Freising-Weihenstephan Germany [email protected]

Volker Liebetrau Leibniz Institute of Marine Sciences IFM-GEOMAR Research Division 2: Marine Biogeochemistry Wischhofstr. 1-3 24148 Kiel Germany [email protected] Wolfgang Liebl Department of Microbiology Technische Universität München Emil-Ramann-Str. 4 85354 Freising-Weihenstephan Germany [email protected]

Kathrin I. Mohr Albrecht-von-Haller-Institute for Plant Sciences University of Göttingen Untere Karspüle 2 37073 Göttingen Germany and Helmholtz Centre for Infection Research Braunschweig Germany [email protected] D. Kirk Nordstrom U.S. Geological Survey 3215 Marine St. Boulder, CO 80303 USA [email protected]

Carol D. Litchfield Department of Environmental Science & Policy George Mason University 10900 University Boulevard Manassas, VA 20110 USA [email protected]

Ronald S. Oremland Water Resources Division, MS 480 United States Geological Survey 345 Middlefield Road Menlo Park, CA 94025 USA [email protected]

Frank E. Löffler Department of Microbiology and Department of Civil and Environmental Engineering University of Tennessee Knoxville, TN 37996 USA [email protected]

Wolfgang Oschmann Geologie and Paläontologie University of Frankfurt Senckenberganlage 32 60325 Frankfurt Germany [email protected]

Fred T. Mackenzie Department of Oceanography School of Ocean and Earth Science and Technology University of Hawaii Honolulu, HI 96822 USA [email protected]

Muriel Pacton Geological Institute ETH Zurich Sonneggstrasse 5 8092 Zurich Switzerland [email protected]

CONTRIBUTORS

Karsten Pedersen Department of Cell and Molecular Microbiology Göteborg University Box 462 405 30 Göteborg Sweden [email protected] Randall S. Perry Department of Earth Science and Engineering Imperial College London UK [email protected] Steven T. Petsch Department of Geosciences University of Massachusetts Amherst Amherst, MA 01003 USA [email protected] Thomas Pommier Microbial Ecology Centre UMR 5557 CNRS-Université Lyon 1; USC 1193 INRA CNRS-Université lyon 43 bd du 11 November 1918 69622 Villeurbanne France [email protected] Hubertus Porada Geoscience Centre University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Mihály Pósfai Department of Earth and Environmental Sciences University of Pannonia P.O. Box 158 8200 Veszprém Hungary [email protected] Nicole R. Posth Geomicrobiology Group Center for Applied Geoscience University of Tübingen Sigwartstrasse 10 72072 Tübingen Germany [email protected]

Brian R. Pratt Department of Geological Sciences University of Saskatchewan Saskatoon, SK S7N 5E2 Canada [email protected] Nadia-Valérie Quéric Geobiology Group Geoscience Center University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Gudrun Radtke Hessisches Landesamt für Umwelt und Geologie Rheingaustr. 186 65203 Wiesbaden Germany [email protected] Eugenio Ragazzi Department of Pharmacology and Anaesthesiology University of Padova 35131 Padova Italy [email protected] Kristina Rathsack Geobiology Group Geoscience Center University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Douglas Eric Rawlings Department of Microbiology University of Stellenbosch Private Bag X1 Matieland 7602 South Africa [email protected] R. Pamela Reid Marine Geology and Geophysics – RSMAS University of Miami 4600 Rickenbacker Causeway Miami, FL 33149 USA [email protected]

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CONTRIBUTORS

Andreas Reimer Geobiology Group Geoscience Center University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Joachim Reitner Geobiology Group Geoscience Center University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Robin W. Renaut Department of Geological Sciences University of Saskatchewan Saskatoon, SK S7N 5E2 Canada [email protected] Anna-Louise Reysenbach Department of Biology Portland State University 246 Science Building 2 1719 SW 10th Avenue Portland, OR 97207 USA [email protected] Robert Riding Department of Earth and Planetary Sciences University of Tennessee 1412 Circle Drive Knoxville, TN 37996 USA [email protected]

Jennifer A. Roberts Department of Geology University of Kansas Lawrence, KS 66045-7613 USA [email protected] Manuel Rodriguez-Gallego Departamento de Mineralogía y Petrología Universidad de Granada Fuentenueva s/n 18002 Granada Spain [email protected] Marta Rodríguez-Martínez Departamento de Geología Facultad de Biología Universidad de Alcalá 28871 Alcalá de Henares, Madrid Spain [email protected] Carlos Rodriguez-Navarro Departamento de Mineralogía y Petrología Universidad de Granada Fuentenueva s/n 18002 Granada Spain [email protected] Michael Rothballer Department Microbe-Plant Interactions Helmholtz Zentrum München German Research Center for Environmental Health (GmbH) 85764 Neuherberg Germany [email protected]

Kathrin Riedel Department of Microbiology Institute of Plant Biology University of Zurich Winterthurerstrasse 190 8057 Zurich Switzerland [email protected]

Michael J. Russell Planetary Science & Life Detection Jet Propulsion Laboratory California Institute of Technology 4800 Oak Grove Drive Pasadena, CA 91109-8099 USA [email protected]

Elizaveta M. Rivkina Institute of Physicochemical & Biological Problems in Soil Science Russian Academy of Sciences Pushchino Russia [email protected]

Jürgen Schieber Department of Geological Sciences Indiana University 1001 E 10th Str Bloomington, IN 47405-1405 USA [email protected]

CONTRIBUTORS

Bernhard Schink Department of Biology University of Konstanz Konstanz Germany [email protected] Marie-Lise Schläppy Max Planck Institute for Marine Microbiology Bremen Germany [email protected] Michael Schmid Department Microbe-Plant Interactions Helmholtz Zentrum München German Research Center for Environmental Health (GmbH) 85764 Neuherberg Germany [email protected] Alexander R. Schmidt Courant Research, Centre Geobiology University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Kerstin Schmidt Institut für Ökologie Friedrich-Schiller-Universität Jena Dornburger Str. 159 07743 Jena Germany [email protected]

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Mark A. Sephton Department of Earth Science and Engineering Imperial College London London SW7 2AZ UK [email protected] Silke Severmann Institute of Marine and Coastal Sciences and Department of Earth and Planetary Sciences Rutgers University 71 Dudley Road New Brunswick, NJ USA [email protected] Meinhard Simon Institute for Chemistry and Biology of the Marine Environment University of Oldenburg 26111 Oldenburg Germany [email protected] Peter Sjövall SP Technical Research Institute of Sweden Chemistry and Materials Technology Borås Sweden [email protected] Helga Stan-Lotter Division of Molecular Biology Department of Microbiology University of Salzburg Billrothstr. 11 5020 Salzburg Austria [email protected]

Carsten J. Schubert Eawag, Swiss Federal Institute of Aquatic Science and Technology Seestrasse 79 6047 Kastanienbaum Switzerland [email protected]

John F. Stolz Bayer School of Natural & Environmental Sciences Department of Biological Sciences Duquesne University 600 Forbes Avenue Pittsburgh, PA 15282 USA [email protected]

Heide N. Schulz-Vogt Max Planck Institute for Marine Microbiology Celsiusstr. 1 28359 Bremen Germany [email protected]

Kristina L. Straub Biogeochemie/Umweltgeowissenschaften University of Vienna Althanstr. 14 1090 Vienna Austria [email protected]

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CONTRIBUTORS

Volker Thiel Geobiology Group Geoscience Center University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Ingunn H. Thorseth Centre of Geobiology and Department of Earth Science University of Bergen Allegaten 41 5007 Bergen Norway [email protected] Jan Toporski Wissenschaftliche Instrumente und Technologie GmbH Lise-Meitner-Strasse 6 89081 Ulm Germany [email protected] Aline Tribollet Institut de Recherche pour le Développement BP A5 98848 Nouméa, Nouvelle-Calédonie France [email protected] Eric P. Verrecchia Institut de Géologie et Paléontologie Université de Lausanne Anthropole 1015 Lausanne Switzerland [email protected] Pieter T. Visscher Center for Integrative Geosciences – Marine Sciences University of Connecticut 354 Mansfield Road Storrs, CT 06269-2045 USA [email protected] Walter Vortisch Prospektion und Angewandte Sedimentologie Department für Angewandte Geowissenschaften und Geophysik Montanuniversität Leoben Leoben Austria [email protected]

Lesley A. Warren School of Geography and Earth Sciences McMaster University Hamilton, ON Canada [email protected] Bettina Weber Plant Ecology and Systematics Department of Biology University of Kaiserslautern P.O. Box 3049 67653 Kaiserslautern Germany [email protected] Frances Westall Centre de Biophysique Moléculaire CNRS Université de Orléans Rue Charles Sadron 45071 Orléans France [email protected] Frank Wiese Courant Research Centre Geobiology University of Göttingen Goldschmidtstr. 3 37077 Göttingen Germany [email protected] Gert Wörheide Department of Earth & Environmental Sciences Palaeontology and Geobiology Ludwig-Maximilians-Universität München Richard-Wagner-Str. 10 80333 München Germany [email protected] Christoph Wrede Institut für Mikrobiologie und Genetik University of Göttingen Göttingen Germany [email protected] Li Zhang Faculty of Earth Sciences China University of Geosciences Wuhan, Hubei 430074 China [email protected]

CONTRIBUTORS

Maoyan Zhu State Key Laboratory of Palaeobiology and Stratigraphy Chinese Academy of Sciences 210008 Nanjing China [email protected]

Frank Zielinski Department of Environmental Microbiology Biotrophic Plant-Microbe Interactions Helmholtz Centre for Environmental Research (UFZ) Permoserstr. 15 04318 Leipzig Germany [email protected]

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Preface

Geobiology is a highly cross-disciplinary field that explores the present and past relationships that life has with non-living matter. “Biosphere meets Geosphere” perhaps most parsimoniously describes the fundamental concept of Geobiology. In 1991, Peter Westbroek, a Dutch paleontologist and influential protagonist of Geobiology defined the field in a book entitled “Life as a Geological Force: Dynamics of the Earth”, thus motivating a new way of thinking in the geosciences. His fundamental work on processes of biomineralization in coccolithophorid algae (Westbroek and de Jong, 1983) greatly contributed to the understanding of metabolic processes controlling mineral formation. Westbroek’s thinking was influenced by James Lovelock’s Gaia concept (Lovelock, 1988) which advocated the importance of biological processes with regard to global change over time. Other early pioneers of the Geobiology concept were the Russian scientist Georgy Adamovich Nadson (1903), who recognised microorganisms as geological agents, and the Swiss geologists Johannes Neher and Ernst Rohrer who discovered the role of microbes in dolomite formation (Neher and Rohrer 1958) and their presence in the deep biosphere of crystalline rocks (Neher and Rohrer 1959). In 1971, the German geoscientist Gerd Lüttig introduced a new discipline that merged aspects of geology and biology and called it “Lithobiontik”. This was the first time that research on geological and biological interactions received a well-founded definition: “Die Erdgeschichte ist umschreibbar als eine ständige Auseinandersetzung zwischen Gesteinswelt (lithos) und Lebewelt (bios). Die Gesamtheit der entsprechenden Vorgänge zu erforschen, ist Aufgabe der Lithobiontik, einer Forschungsrichtung im Grenzgebiet zwischen Geologie und Biologie.” — Earth history can be described as a permanent interaction between the geosphere (lithos) and life processes (bios). To investigate these processes is the mission of Lithobiontics, a new research discipline between Geology and Biology.

Kenneth Nealson and William Ghiorse collegues provided, in a conceptual review written for the American Academy of Microbiology (Nealson et al. 2001), a modern and concise perception of Geobiology as “Exploring the interface between the Biosphere and the Geosphere”. The interplay between biological and geological processes has shaped the Earth and driven the evolution of its biodiversity from the early dawn of life, some four billion years ago. Since then, organisms have been responding to a changing global environment and in turn, have themselves altered the chemical and physical settings on our planet. Geobiology strives to identify these causeand-effect chains in both modern environments and the geological record. Its goal is to provide, on different time and spatial scales, an ‘organismic’ biological perspective on Earth’s environmental evolution (Knoll and Hayes, 1997). Shifting their focus from traditional morphological studies made by paleontologists, geobiologists continue to develop their ability to relevant chemical and molecular signatures within living and non-living materials, and to interpret these signatures to better understand the geological record and better predict our course into the future. A key issue in most geobiological studies is the elucidation of ancient environmental states, and to understand the evolution of biological processes and their geological consequences. Meaningful signatures that reveal such processes are not limited to visible remains, but also encompass for instance, organic molecules, minerals, petrofabrics and isotope patterns. Organisms may alter their chemical environment, thereby giving rise to the production or destruction of minerals, rocks, atmospheric gases and even fossil fuels. An understanding of these processes creates enormous potential with respect to issues of environment protection, public health, energy and resource management. Geobiological research and

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education is therefore rapidly growing, with topics becoming more common in University and High School curricula. With Geobiology issues including many spectacular aspects (e.g., early life, deep biosphere, gas hydrates, black smokers etc.), public interest also increases concomittantly. Moving beyond the borders of classical core disciplines, scientists from a broad range of disciplines are actively involved in geobiological studies. There is no common perception of a ‘typical’ geobiologist, but as an underlying requirement, scientists need to adapt concepts and utilize methodologies from other disciplines to exploit the full potential of Geobiology. Fields united under the umbrella of geobiology include, but are not limited to: geology and paleontology, mineralogy, microbiology, molecular biology, genomics, organic and inorganic geochemistry, oceanography, astrobiology and (paleo)ecology. This incomplete list provides an impression of the implications of geobiological research, and that the super-discipline of Geobiology is ‘greater than the sum of its parts’ (Nealson et al. 2001). The Encyclopedia of Geobiology was compiled to provide clear explanations of current geobiological topics. It is not structured as a student textbook, but rather to quickly access particular terms and concepts in self-contained entries. We hope that this volume will also tempt the casual reader to browse and become curious about the different facettes and foci of Geobiology - following the philosophy of the late Founding Series Editor, Rhodes Fairbridge, we will be most content if an original discovery may emerge from perusing a variety of entries. We are thankful to our authors, both distinguished ‘old hands’ and young researchers from five continents, who have synthesized the particular subdisciplines for this volume. The preparation of this book was greatly aided by the input we received from our Editorial Board members Hans-Joachim Fritz, Andreas Kappler, Kurt Konhauser,

Pamela Reid, and Xingliang Zhang, and we wish to express our sincere thanks to them for their invaluable support and encouragement. Göttingen, October 2010 The Editors-In-Chief Joachim Reitner Volker Thiel

References Knoll, A. H., and Hayes, J. M., 2000. Geobiology: problems and prospects. In Lane, R. H., Steininger, F. F., Kaesler, R. L., Ziegler, W., Lipps, J. (eds.), Fossils and the Future - Paleontology in the 21st Century. Senckenberg-Buch Nr. 74, Frankfurt a. M.: Kramer, pp. 149–153. Lovelook, J. E., 1988. The Ages of Gaia. New York: Norten. Lüttig, G., 1971. Lithobiontik – Aufgabengebiet, Tätigkeiten, zukünftige Ziele (Lithobiontics – scope, activities, future goals). Geol Jb (in German), 89, 575–582. Nadson, G. A., 1903. Microorganismi kak geologitsheskie dieiatieli (Microorganisms as geological agents). I. Tr. Komisii Isslect. Min. Vodg. Slavyanska (in Russian), St. Petersburg, 1–98. Nealson, K., Ghiorse, W. A., Strauss, E., 2001. Geobiology: Exploring the Interface Between the Biosphere and the Geosphere. Washington, D.C., American Academy of Microbiology, 16 p.; available online at: http://academy.asm.org/index.php/ colloquia-reports/browse-all/227-geobiology-exploring-theinterface-between-the-biosphere-and-the-geophete-2001-b. Neher, J., and Rohrer, E., 1958. Dolomitbildung unter Mitwirkung von Bakterien (Dolomite formation involving bacteria). Eclogae Geol Helv (in German), 51, 213–215. Neher, J., and Rohrer, E., 1959. Bakterien in tieferliegenden Gesteinslagen (Bacteria in deeper rock layers). Eclogae Geol Helv (in German), 52, 619–625. Westbroek, P., 1991. Life as a Geological Force – Dynamics of the Earth. New York: W.W.N. Norten. Westbroek, P., 1983. Biological metal accumulation and biomineralization in a geological perspectives. In Westbroek P. and de Jong, W. E. (eds.), Biological Metal Accumulation and Biomineralization in a Geological Perspective, Dordrecht: Reidel, pp. 1–11.

A

ACETOGENS Kirsten Küsel1, Harold L. Drake2 1 Friedrich Schiller University Jena, Jena, Germany 2 University of Bayreuth, Bayreuth, Germany

Synonyms Homoacetogens Definition Acetogens are defined as anaerobic prokaryotes that use the acetyl-CoA pathway for the (a) reductive synthesis of the acetyl moiety of acetyl-CoA from CO2, (b) conservation of energy, and (c) assimilation of CO2 into biomass. Introduction Acetogens utilize the acetyl-CoA “Wood–Ljungdahl” pathway as a terminal electron-accepting, energy-conserving, CO2-fixing process. The reductive synthesis of acetate from CO2 differentiates acetogens from organisms that synthesize acetate by other metabolic processes. Although the production of acetate as a sole reduced end product is the classic hallmark of acetogens, the production of acetate is not a part of the definition, because the acetogen might not form acetate in situ or when cultured in the laboratory. The first acetogen, Clostridium aceticum, was isolated from soil by the Dutch microbiologist K. T. Wieringa in 1936. This spore-forming, mesophilic bacterium was shown to grow at the expense of H2–CO2 and synthesizes acetate according to the following stoichiometry (Wieringa, 1939–1940): 4H2 þ 2CO2 ! CH3 COOH þ 2H2 O In 1936, this reaction constituted a unique mechanism for the fixation of CO2. Unfortunately, the culture of

C. aceticum was lost and no further work was done until it was re-isolated in the early 1980s (Braun et al., 1981). In 1942, F. E. Fontaine and coworkers isolated the second acetogen, Clostridium thermoaceticum (later reclassified as Moorella thermoacetica), a spore-forming, thermophilic bacterium that catalyzed the near stoichiometric conversion of glucose to acetate (Fontaine et al., 1942): C6 H12 O6 ! 3CH3 COOH C. thermoaceticum became the most extensively studied acetogen and was used to resolve the enzymology of the acetyl-CoA pathway in the laboratories of two biochemists, H. G. Wood and L. G. Ljungdahl (Drake and Daniel, 2004). The demonstration that 14CO2 was incorporated equally into both carbons of acetate by C. thermoaceticum was the first 14C-study in biology (Barker and Kamen, 1945). The synthesis of acetate from two molecules of CO2 was later confirmed by Wood using 13CO2 and mass spectrometry (Wood, 1952). Both studies demonstrated that acetogens featured a new autotrophic mechanism for the fixation of CO2. The acetyl-CoA pathway, also referred to as the Wood–Ljungdahl pathway, is now recognized as a fundamental component of the global carbon cycle, important to primary production in subsurfaceecosystems, and essential to the metabolic potentials of not only acetogens but also methanogens, sulfatereducers, and anammox bacteria (Wood and Ljungdahl, 1991; Drake, 1994; Schouten et al., 2004).

The acetyl-CoA Wood–Ljungdahl pathway The acetyl-CoA pathway is a reductive, linear, “onecarbon” process (Figure 1) in contrast to cyclic CO2-fixing processes (Calvin cycle, reductive tricarboxylic acid cycle, and hydroxypropionate cycle). The two branches of the acetyl-CoA pathway merge at the synthesis of acetyl-CoA that is converted to either acetate or

Joachim Reitner & Volker Thiel (eds.), Encyclopedia of Geobiology, DOI 10.1007/978-1-4020-9212-1, # Springer Science+Business Media B.V. 2011

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2002; Russel and Martin, 2004). It has been proposed that biochemistry started when marine CO2 from volcanoes and hydrothermal H2 met at a hydrothermal vent rich in metal sulfides, where an analogue of the exergonic acetyl-CoA pathway catalyzed the synthesis of organic precursors to fuel primordial biochemical reactions.

Acetogens, Figure 1 The acetyl-CoA “Wood-Ljungdahl” pathway. H4F, tetrahydrofolate; HSCoA, coenzyme A; Pi, inorganic phosphate; e, electron; co-protein, corrinoid enzyme; ATP, adenosine-5’-triphosphate. The pathway is from Mu¨ller et al. (2004) and used with kind permission of Horizon Bioscience.

assimilated into biomass (Drake, 1994; Drake et al., 2006). Acetyl-CoA synthase (ACS) not only catalyzes the reduction of CO2 to CO and the synthesis of acetyl-CoA but also oxidizes CO to CO2. Thus, the acetyl-CoA synthase is also termed CO dehydrogenase (CODH). Energy conservation occurs during the reductive synthesis of acetate by substrate-level phosphorylation and chemiosmotic processes. For each glucose that is converted to three acetates, four molecules of ATP are produced by substrate-level phosphorylation (ATPSLP). However, when growth occurs under autotrophic conditions, there is no net gain in ATPSLP, and growth is dependent upon translocation of protons or sodium ions. The relatively simple features of the linear acetyl-CoA pathway and the fact that metals and metal sulfides do the biochemical work of CO2 fixation in the key enzymes of the pathway (i.e., CODH and ACS) suggest that the acetyl-CoA pathway might have been the first autotrophic and early respiratory process on earth and, thus, important in the evolution of life (Wood, 1991; Miyakawa et al.,

Phylogenetic diversity of acetogens Over 100 acetogenic bacterial species have been isolated to date from very diverse habitats, including marine and freshwater sediments; salt lake soda deposits; tundra, forest, and agricultural soils; the subsurface; fecal material; and marine and salt marsh plants (Drake et al., 2006, 2008). Acetogens display extreme genetic diversity, having genomic G + C contents that vary between 22 mol% (Clostridium ljungdahli) to 62 mol% (Holophaga foetida). Acetogens have been assigned to 22 different bacterial genera including Acetitomaculum, Acetoanaerobium, Acetobacterium, Acetohalobium, Acetonema, Bryantella, “Butyribacterium,” Caloramator, Clostridium, Eubacterium, Holophaga, Moorella, Natroniella, Natronincola, Oxobacter, Ruminococcus, Sporomusa, Syntrophococcus, Tindallia, Thermoacetogenium, Thermoanaerobacter, and Treponema (the name in quotation marks has not been validated). The diverse habitat range of acetogens demonstrates that they are adapted to a broad range of in situ conditions. Many acetogens are sporeformers (e.g., Byrer et al., 2000), a feature that aids in surviving unfavorable in situ conditions, and some acetogens have connecting filaments (Küsel et al., 2001) that might facilitate communication. The usage of the acetyl-CoA pathway for autotrophic assimilation of carbon and acetate utilization by methanogens suggests that Archaea also can grow via acetogenesis. Indeed, the methanogen Methanosarcina acetivorans C2A uses the acetyl-CoA pathway to convert carbon monoxide to both acetate and methane, that is, methane is not the sole reduced end product of this methanogen (Rother and Metcalf, 2004; Lessner et al., 2006). Similarly, the archaea Archaeoglobus fulgidus VC16 can grow via CO-dependent acetogenesis (Henstra et al., 2008). Only some acetogenic genera, like Moorella and Sporomusa, are monophyletic, but many acetogens are phylogenetically dispersed within genera that contain non-acetogenic species. Thus, the phylogenetic position of 16S rRNA gene sequences is usually inadequate for resolving the functional identity of a potential acetogen. To date, only some highly specific 16S rRNA-based probes and primers have been designed to target subsets of acetogenic taxa (Küsel et al., 1999). Molecular approaches based on the analysis of functional genes central to the acetyl-CoA pathway (e.g., a gene for formyltetrahydrofolate synthetase) are still limited due to problems of probe specificity (Leapheart et al., 2003). Functional diversity of acetogens Acetogens utilize a wide variety of electron donors and electron acceptors and can engage alternative terminal

ACETOGENS

3

electron-accepting processes when challenged with O2 (Drake et al., 2006). Many acetogens can utilize one or more terminal electron-accepting processes in addition to acetogenesis. For example, nitrate is a preferred electron acceptor for M. thermoacetica and is dissimilated to nitrite and ammonium (Seifritz et al., 1993), and perchlorate is a preferred terminal electron acceptor for Moorella perchloratireducens (Balk et al., 2008). The engagement of diverse redox couples enables acetogens to form junction points within and between biological cycles at the ecosystem level (Drake and Küsel, 2005). Acetogens are known to use the following alternative terminal electron acceptors that yield diverse end products listed below: Fumarate ! succinate Methoxylated phenylacrylates ! methoxylated phenyl propionates Nitrate ! nitrite Nitrite ! ammonium Thiosulfate ! sulfide Dimethylsulfoxide ! dimethylsulfide Perchlorate ! chloride Chlorate ! chloride Pyruvate ! lactate Acetaldehyde ! ethanol Protons ! hydrogen gas CO, H2, carbohydrates, alcohols, carboxylic acids, dicarboxylic acids, aldehydes, substituent groups of various aromatic compounds, and certain halogenated substrates are examples of substrates that acetogens can oxidize. Although hexoses or pentoses are utilized by most of the acetogens isolated to date, none of these isolates appear to be able to degrade high molecular weight polymers, such as cellulose and lignin. Most acetogens have the capacity to produce more than acetate as their sole reduced end product, and this capacity is dependent on the availability of both reductant and terminal electron acceptors, including CO2. Thus, referring to acetogens as homoacetogens is usually a misnomer. In anoxic habitats, acetogens compete with primary fermentors for monomeric compounds that are derived from the initial breakdown of cellulose and lignin and with secondary fermentors for fermentation products. Thermodynamic considerations suggest that acetogenesis should not be a highly competitive microbial process. Nonetheless, acetogens can outcompete methanogens for H2 in freshwater sediments with low pH and low temperature and in the hindgut of certain termites (Conrad et al., 1989; Breznak and Kane, 1990; Drake et al., 2006). Microbially produced acetate can provide up to 100% of the energy requirement of wood-feeding termites. The attachment of H2-consuming acetogenic spirochetes to H2-producing protozoa provides H2 concentrations above the known H2-threshold values for acetogens. Another ecological advantage might be the low oxygen sensitivity of certain acetogens (Figure 2). Acetogens have been classically referred to as obligate, if not strict, anaerobes, and many enzymes central to acetogenesis are

Commensal consumption of O2 Microaerophile or aerotolerant fermenter

Acetogen

O2 Carbohydrate

X

Acetate

H2O

1

Oxidative stress

2

Reductive detoxification O2

3

Metabolic switch to fermentation e–

O2.–

H2O2

CO2 e–

H2O

e–

Pyruvate

e–

H2O

H2O2

Oxic conditions Acetate Lactate

Acetogens, Figure 2 Mechanisms by which acetogens cope with oxidative stress. X, products (e.g., H2, formate, lactate) that are derived from the partial oxidation of carbohydrates (in some cases, short-chain polymers [e.g., stachyose] that are not substrates for the acetogen); e, electron. (Modified from Mu¨ller et al., 2004, and used with kind permission from Horizon Bioscience.)

extremely sensitive to O2. However, acetogens have been isolated from oxic habitats, like soils and the rhizosphere of macrophytes that release O2 through their roots, indicating that such acetogenic species must cope with periods of oxidative stress. Acetogens contain numerous enzymes that can reductively remove O2 and its toxic by-products (e.g., superoxide and peroxide), when the concentration of O2 is relatively low (Drake et al., 2006). These enzymes include peroxidase, NADH-oxidase, rubredoxin oxidoreductase (a superoxide reductase), rubrerythrin (a peroxidase), superoxide dismutase, catalase, and cytochrome bd oxidase (Das et al., 2001, 2005; Küsel et al., 2001). Acetogens can shift the flow of reductant away from the acetyl-CoA pathway to alternative terminal electron-accepting processes that are less sensitive to O2 and operate at higher redox potentials than the very low standard redox potential of the CO2/acetate half-cell reaction (290 mV). For example, Clostridium glycolicum RD-1 (isolated from seagrass roots) is an aerotolerant acetogen that switches from acetogenesis to

4

ACETOGENS

classic fermentation in response to O2 (Küsel et al., 2001). Acetogens can also form symbiotic relationships with O2consuming microaerophiles and aerotolerant fermenters (Gößner et al., 1999). The microaerophile can be a fermentative non-acetogen that has the capacity to consume O2 and forms fermentation products (e.g., lactate, formate, and H2) that can be used by the acetogen for acetogenesis. Such interactions can protect acetogens from oxidative stress and form trophic linkages in habitats with fluctuating redox conditions.

Conclusions It has been estimated that approximately 1012 kg of acetate is synthesized per year via H2-driven acetogenesis in sediments and the hindgut of termites (Breznak and Kane, 1990), a number that is fivefold greater than the annual amount of methane produced via the methanogenic reduction of CO2. Such estimates accentuate the potential importance of acetogens and acetogenesis to the global carbon cycle. However, in situ acetate turnover measurements are complicated and acetogens catalyze a large number of redox reactions, suggesting that their in situ activities are not restricted to acetogenesis. Thus, although acetate forms an important trophic link in a wide variety of ecosystems, in situ information on acetogens and their activities are often theoretical. Bibliography Andreesen, J. R., and Ljungdahl, L. G., 1973. Formate dehydrogenase of Clostridium thermoaceticum: incorporation of selenium75, and the effects of selenite, molybdate, and tungstate on the enzyme. The Journal of Bacteriology, 116, 867–873. Balk, M., van Gelder, T., Weelink, S. A., and Stams, A. J. M., 2008. (Per)chlorate reduction by the thermophilic bacterium Moorella perchloratireducens sp. nov., isolated from underground gas storage. Applied and Environmental Microbiology, 74, 403–409. Barker, H. A., and Kamen, M. D., 1945. Carbon dioxide utilization in the synthesis of acetic acid by Clostridium thermoaceticum. Proceedings of the National Academy of Sciences of the United States of America, 31, 219–225. Braun, M., Mayer, F., and Gottschalk, G., 1981. Clostridium aceticum (Wieringa), a microorganism producing acetic acid from molecular hydrogen and carbon dioxide. Archives of Microbiology, 128, 288–293. Breznak, J. A., and Kane, M. D., 1990. Microbial H2/CO2 acetogenesis in animal guts: nature and nutritional significance. FEMS Microbiology Reviews, 7, 309–313. Byrer, D. E., Rainey, F. A., and Wiegel, J., 2000. Novel strains of Moorella thermoacetica form unusually heat-resistant spores. Archives of Microbiology, 174, 334–339. Conrad, R., Bak, F., Seitz, H. J., Thebrath, B., Mayer, H. P., and Schütz, H., 1989. Hydrogen turnover by psychrotrophic homoacetogenic and mesophilic methanogenic bacteria in anoxic paddy soil and lake sediment. FEMS Microbiology Ecology, 62, 285–294. Das, A., Coulter, E. D., Kurtz, D. M., and Ljungdahl, L. G., 2001. Five-gene cluster in Clostridium thermoaceticum consisting of two divergent operons encoding rubredoxin oxidoreductaserubredoxin and rubrerythrin-type A flavoprotein-highmolecular-weight rubredoxin. The Journal of Bacteriology, 183, 1560–1567.

Das, A., Silaghi-Dumitrescu, R., Ljungdahl, L. G., and Kurtz, D. M., Jr., 2005. Cytochrome bd oxidase, oxidative stress, and dioxygen tolerance of the strictly anaerobic bacterium Moorella thermoacetica. The Journal of Bacteriology, 187, 2020–2029. Drake, H. L. 1994. Acetogenesis, acetogenic bacteria, and the acetyl-CoA “Wood/Ljungdahl” pathway: past and current perspectives. In Drake, H. L. (ed.), Acetogenesis. New York: Chapman & Hall, pp. 3–60. Drake, H. L., and Daniel, S. L., 2004. Physiology of the thermophilic acetogen Moorella thermoacetica. Research in Microbiology, 155, 869–883. Drake, H. L., and Küsel, K., 2005. Acetogenic clostridia. In Dürre, P. (ed.), Handbook on Clostridia. Boca Raton, FL: CRC Press, pp. 719–746. Drake, H. L., Küsel, K., and Matthies, C., 2006. Acetogenic prokaryotes. In Dworkin, M., Falkow, S., Rosenberg, E., Schleifer, K.-H., and Stackebrandt, E. (eds.), The Prokaryotes, Vol. 2. New York: Springer-Verlag, pp. 354–420. Drake, H. L., Gößner, A. S., and Daniel, S. L., 2008. Old acetogens, new light. Annals New York Academy of Sciences, 1125, 100–128. Fontaine, F. E., Peterson, W. H., McCoy, E., and Johnson, M. J., 1942. A new type of glucose fermentation by Clostridium thermoaceticum n. sp. The Journal of Bacteriology, 43, 701–715. Gößner, A. S., Devereux, R., Ohnemüller, N., Acker, G., Stackebrandt, E., and Drake, H. L., 1999. Thermicanus aegyptius gen. nov., sp. nov., isolated from oxic soil, a fermentative microaerophile that grows commensally with the thermophilic acetogen Moorella thermoacetica. Applied and Environmental Microbiology, 65, 5124–5133. Henstra, A. M., Dijkema, C., and Stams, A. J. M., 2008. Archaeoglobus fulgidus couples CO oxidation to sulfate reduction and acetogenesis with transient formate accumulation. Environmental Microbiology, 9, 1836–1841. Küsel, K., Pinkart, H. C., Drake, H. L., and Devereux, R., 1999. Acetogenic and sulfate-reducing bacteria inhabiting the rhizoplane and deeper cortex cells of the sea grass Halodule wrightii. Applied and Environmental Microbiology, 65, 5117–5123. Küsel, K., Karnholz, A., Trinkwalter, T., Devereux, R., Acker, G., and Drake, H. L., 2001. Physiological ecology of Clostridium glycolicum RD-1, an aerotolerant acetogen isolated from sea grass roots. Applied and Environmental Microbiology, 67, 4734–4741. Leaphart, A. B., Friez, M. J., and Lovell, C. R., 2003. Formyltetrahydrofolate synthetase sequences from salt marsh plant roots reveal a diversity of acetogenic bacteria and other bacterial functional groups. Applied and Environmental Microbiology, 69, 693–696. Lessner, D. J., Li, L., Li, Q., Rejtar, T., Andreev, V. P., Reichlen, M., Hill, K., Moran, J. J., Karger, B. L., and Ferry, J. G. 2006. An unconventional pathway for reduction of CO2 to methane in CO-grown Methanosarcina acetivorans revealed by proteomics. Proceedings of the National Academy of Science of the United States of America, 103, 17921–17926. Ljungdahl, L. G., 1986. The autotrophic pathway of acetate synthesis in acetogenic bacteria. Annual Reviews of Microbiology, 40, 415–450. Miyakawa, S., Yamanashi, H., Kobayashi, K., Cleaves, H. J., and Miller, S. L., 2002. Prebiotic synthesis from CO atmospheres: implications for the origins of life. Proceedings of the National Academy of Science of the United States of America, 99, 14628–14631. Müller, V., Inkamp, F., Rauwolf, A., Küsel, K., and Drake, H. L., 2004. Molecular and cellular biology of acetogenic bacteria. In Nakano, M., and Zuber, P. (eds.), Strict and Facultative Anaerobes: Medical and Environmental Aspects. Norfolk, UK: Horizon Scientific Press, pp. 251–281.

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Rother, M., and Metcalf, W. W., 2004. Anaerobic growth of Methanosarcina acetivorans C2A on carbon monoxide: an unusual way of life for a methanogenic archaeon. Proceedings of the National Academy of Science of the United States of America, 101, 16929–16934. Russell, M. J., and Martin, W., 2004. The rocky roots of the acetyl-CoA pathway. Trends in Biochemical Sciences, 29, 358–363. Schouten, S., Strous, M., Kuypers, M. M. M., Irene, W., Rijpstra, C., Baas, M., Schubert, C. J., Jetten, M. S. M., and Damste, J. S. S., 2004. Stable carbon isotopic fractionations associated with inorganic carbon fixation by anaerobic ammonium-oxidizing bacteria. Applied and Environmental Microbiology, 70, 3785–3788. Seifritz, C., Daniel, S. L., Gößner, A., and Drake, H. L., 1993. Nitrate as a preferred electron sink for the acetogen Clostridium thermoaceticum. The Journal of Bacteriology, 175, 8008–8013. Wieringa, K. T., 1939–1940. The Formation of acetic acid from carbon dioxide and hydrogen by anaerobic spore-forming bacteria. Antonie Van Leeuwenhoek Journal of Microbiology and Serology, 6, 251–262. Wood, H. G. 1952. A study of carbon dioxide fixation by mass determination on the types of C13-acetate. Journal of Biological Chemistry, 194, 905–931. Wood, H. G. 1991. Life with CO or CO2 and H2 as a source of carbon and energy. Journal of the Federation of American Societies for Experimental Biology, 5, 156–163. Wood, H. G., and Ljungdahl, L. G., 1991. Autotrophic character of acetogenic bacteria. In Shively, J. M., and Barton, L. L. (eds.), Variations in Autotrophic Life. San Diego, CA: Academic, pp. 201–250.

Cross-references Anaerobic Transformation Processes, Microbiology Bacteria Carbon (Organic, Cycling) Fermentation Hydrogen Methanogens Microbial Communities, Structure, and Function Microbial Degradation Origin of Life

ACID ROCK DRAINAGE Lesley A. Warren McMaster University, Hamilton, ON, Canada

Synonyms Acid mine drainage; AMD; ARD Definition Acid mine and acid rock drainage (AMD/ARD) refer to the extremely acidic (pH < 3), metal-rich waters that are derived from the weathering of sulfidic minerals when exposed to air, water, and microorganisms (Figure 1; Nordstrom and Alpers, 1999; Bond et al., 2000):

Acid Rock Drainage, Figure 1 An AMD stream at a nickelcopper mine in northern Ontario, Canada. The orange and red colours in the stream are Fe-oxyhydroxide and Fe-sulphate mineral precipitates.

Environmental significance AMD/ARD is considered to be the most important and widespread global-mining-industry-related pollution problem (Rowe et al., 2007). The main characteristics of ARD/AMD are (1) low pH, (2) high concentrations of dissolved heavy metals, and (3) high concentrations of sulfate (SO4 2, Tsukanoto et al., 2004). Total dissolved metal concentrations as high as 200,000 mg/L and dissolved SO4 2 as high as 760,000 mg/L have been reported to be associated with AMD (Nordstrom et al., 2000). AMD/ARD generation AMD is generated through a combination of chemical and biological (microorganism) processes by which sulfidic minerals are converted to sulfates and iron oxyhydroxides

6

ACID ROCK DRAINAGE

when exposed to water and oxygen (see Sulfide Mineral Oxidation). The overall oxidative dissolution of pyrite (FeS2), a widespread common mine constituent, and associated with AMD generation is given by Equation 1:

At pH values pH 4, iron removed through Fe oxyhydroxide precipitation Fe(OH)3 + 3H+ 3H2O + Fe3+ + 1/4H2O

1/4O2 + 2H+ + 2 Fe2+ + 2SO4 – + 2H+

Reaction continues and pH decreases

pH < 4 Fe 3+ solubility greater at pH < 4 O2 + H2O

FeS 2

Fe3+ +

+ 1/4H2O

FeS2 + 7/2O2 + H2O

FeS2 + 14Fe3+ + 8H2O

1/4O2 + 2H+

+ Fe2+ + 2SO4

2–

+ 2H+

Iron-oxidizing bacteria accelerate Fe oxidation (factors > 10 6)

15Fe2+ + 2SO42– + 16H+

Fe3+-driven pyrite oxidation releases greater acidity than O2

Acid Rock Drainage, Figure 2 A schematic of AMD generation based on FeS2, indicating the relative importance of O2 vs Fe3+-driven oxidation rates and the role of microorganisms as a function of pH. At low pH, Fe3+-driven pyrite oxidation releases greater acidity than O2-driven pyrite oxidation, which is more favourable at circumneutral pH values.

ACID ROCK DRAINAGE

Treatment of acid mine drainage AMD waters must be treated to remove metals and raise the pH before they are discharged to a receiving environment (Neculita et al., 2007). Numerous approaches, often characterized as active or passive, exist to treat AMD. Active processes refer to operations that require a relatively high degree of management and continuous input of consumables, while passive processes theoretically require minimal management and other costs once they are established (Johnson, 2006). The most widespread method is active treatment involving the addition of a chemical neutralizing agent or base (i.e., lime or Ca oxide, CaCO3, Na/Mg carbonates, Na/Mg hydroxides) to increase water pH and precipitate metals as hydroxides and carbonates. Over the last 20 years, there have been considerable new developments for AMD treatment including constructed wetlands (Johnson and Hallberg, 2002), bioreactors, and permeable reactive barriers (Blowes et al., 2000), which take advantage of the increasing knowledge of microbially driven reactions. Much of the biotechnology developments rely on sulfate-reducing bacteria (SRB) to treat AMD. Fundamentally, bioreactors use SRB to drive sulfide precipitation, removing metals from the AMD solution (Equations 6 and 7): 2CH2 O þ SO42 ! 2HCO3 þ H2 S

(6)

H2 S þ MZþ ! MSðsÞ þ 2Hþ

(7)

Z+

where M is a cationic metal such as Cu or Ni. In addition, SRB will drive alkalinity generation (Equation 6) (Koschorreck and Tittel, 2007). However, all emerging treatment approaches are challenged with regard to longterm efficiency. In large part, this reflects the challenges in identifying the complex and dynamic reactions involved in the overall AMD process. The exact mechanisms and controls involved in reactions as well as an understanding and identification of the microbial flora required to efficiently drive desired reactions for bioreactors over the long-term have not yet been elucidated (Edwards et al., 2000; Bernier and Warren, 2005; Bernier and Warren, 2007). In particular, growing evidence of the importance of consortia or mixed microorganism communities in driving AMD-associated processes underscores the need to more fully investigate the interactive roles of bacteria, system pH, [Fe], [O2], and other [metals] involved in the process to narrow the gap in determining effective long-term strategies to mitigate AMD.

Summary AMD/ARD refer to metal-rich acidic waters that are generated through the exposure of sulfidic minerals in mine wastes to water, oxygen, and microorganisms. These discharges are a significant global pollution issue associated with mining activities. Both sulfur- and iron-oxidizing bacteria are involved with Fe-oxidizing bacteria playing a key role in the increased rates of AMD generation compared to abiotic (nonbiologically stimulated oxidation) sulfidic wastes. AMD must be treated for the high levels of acidity and metals before it can be released to the

7

environment. New treatment strategies are emerging that include some combination of chemical and biological approaches; however, all are challenged with regard to long-term efficacy. New research evaluating the roles of environmentally mixed communities of microorganisms demonstrates the opportunities to harness environmentally occurring microorganisms to combat AMD.

Bibliography Berghorn, G. H., and Hunzeker, G. R., 2001. Passive Treatment Alternatives for Remediating Abandoned-Mine Drainage. New York: Wiley, pp. 111–127. Bernier, L., and Warren, L. A., 2005. Acidity generation in a mine tailings lake. Geobiology, 3, 115–133. Bernier, L., and Warren, L. A., 2007. Geochemical diversity in S processes mediated by culture-adapted and environmental enrichments of Acidithiobacillus spp. Geochimica et Cosmochimica Acta, 71, 5684–5697. Blowes, D. W., Ptacek, C. J., Benner, S. G., McRae, C. W. T., Bennett, T. A., and Puls, R. W., 2000. Treatment of inorganic contaminants using permeable reactive barriers. Journal of Contaminant Hydrogeology, 45, 123–137. Bond, P. L., Druschel, G. K., and Banfield, J. K., 2000. Comparison of acid mine drainage microbial communities in physically and geochemically distinct ecosystems. Applied and Environmental Microbiology, 66, 4962–4971. Edwards, K. J., Bond, P. L., Druschel, G. K., McGuire, M. M., Hamers, R. J., and Banfield, J. F., 2000. Geochemical and biological aspects of sulfide mineral dissolution: lessons from iron mountain, California. Chemical Geology, 169, 383–397. Johnson, D. B., 2006. Biohydrometallurgy and the environment: intimate and important interplay. Hydrometallurgy, 83, 153–166. Johnson, D. B., and Hallberg, K. B., 2002. Pitfalls of passive mine water treatment. Reviews in Environmental Sciences and Biotechnology, 1, 335–343. Koschorreck, M., and Tittel, J., 2007. Natural alkalinity generation in neutral lakes affected by acid mine drainage. Journal of Environmental Quality, 36, 1163–1171. Neculita, C.-M., Zagury, G. J., and Bussiere, B., 2007. Passive treatment of acid mine drainage in bioreactors using sulfate-reducing bacteria: critical review and research needs. Journal of Environmental Quality, 36, 1–16. Nordstrom, D. K., and Alpers, C. N., 1999. Geochemistry of acid mine waters. In Plumlee G. S., and Logsdon, M. J. (eds.), The Environmental Geochemistry of Mineral Deposits. Part A: Processes, Techniques, and Health Issues. Littleton, CO: The Society of Economic Geologists, pp. 133–1160. Nordstrom, D. K., and Southam, G., 1997. The geomicrobiology of acid mine drainage. In Banfield, J. F., and Nealson, K. H. (eds.), Geomicrobiology: Interactions Between Microbes and Minerals. Washington, DC: Reviews in Mineralogy Mineralogical Society of America, pp. 361–390. Nordstrom, D. K., Alpers, C. N., Ptacek, C. J., and Blowes, D. W., 2000. Negative pH and extremely acidic mine waters from Iron Mountain, California. Environmental Science and Technology, 34, 254–258. Rowe, O. F., Sanchez-Espana, J., Hallberg, K. B., and Johnson, D. B., 2007. Microbial communities and geochemical dynamics in an extremely acidic, metal-rich stream at an abandoned sulfide mine (Huelva, Spain) underpinned by two functional primary production systems. Environmental Microbiology, 9, 1761–1771. Tsukamoto, T. K., Killion, H. A., and Miller, G. C., 2004. Column experiments for microbiological treatment of acid mine drainage: low temperature, low-pH and matrix investigation. Water Research, 38, 1405–1418.

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ACIDOPHILES

Cross-references Biomining (Mineral Bioleaching, Mineral Biooxidation) Copper Extreme Environments Fe(II)-Oxidizing Prokaryotes Iron Sulfide Formation Ores, Microbial Precipitation and Oxidation Sulfide Mineral Oxidation

ACIDOPHILES “Acidophiles” are organisms thriving in environments below pH 5. For details, see entries “Extreme Environments,” “Biomining (Mineral Bioleaching, Mineral Biooxidation),” “Hot Springs and Geysers,” Hydrothermal Environments (Marine) and “Acid Rock Drainage.”

ACRITARCHS Acritarchs are organic-walled acid-resistant microfossils known from the Proterozoic and throughout the Phanerozoic. The position of these microfossils is still uncertain. A number of acritarch genera have been assigned to Green Algae; others are considered to bear some resemblance to the cysts of dinoflagellates. For more information, please refer to entries “Algae, Eukaryotic” and “Protozoa.”

AEROBIC METABOLISM Heribert Cypionka University of Oldenburg, Oldenburg, Germany

Synonyms Aerobic respiration; Oxygen metabolism Definition Aerobic metabolism comprises the reduction of molecular oxygen as electron acceptor of aerobic respiration, its use as cosubstrate in the degradation of certain compounds, and reactions leading to the detoxification of partially reduced oxygen species. Introduction Life has evolved in the absence of molecular oxygen (see Chapter Early Earth). Therefore, it is not surprising that basic metabolic pathways involved in growth and cell division (like DNA replication or protein synthesis) do not depend on the presence of molecular O2. The evolution of the oxygen-producing (oxygenic) phototrophs dramatically changed the situation of living organisms. Molecular oxygen raised the redox potential of the environment and enabled microorganisms to respire with a much higher energy yield than before. Although oxygen is involved in

a limited number of reactions only, its presence or absence has fundamental impact on biogeochemical processes.

Aerobic degradation of organic matter Aerobic respiration consumes a major part of the photosynthetically produced O2 and about 90% of the organic matter (see Chapter Carbon Cycle). Aerobic (oxygen-metabolizing) organisms normally oxidize their substrates completely to CO2. In most cases, O2 is not directly involved in this process. Substrate oxidation generates reduced electron carriers by means of the respiratory chain. Oxygen is reduced as the last step of the respiration process. Respiratory chain Respiration is a membrane-bound process that generates a chemiosmotic proton gradient across the membrane (Mitchell, 1966). Respiratory chains are composed of series of enzymes, which are organized in multienzyme complexes and use cofactors that allow the stepwise transport of reducing equivalents to oxygen (Figure 1). In eukaryotes (plants, animals, fungi) it takes place within the mitochondria, which are assumed to have evolved from endosymbiotic bacteria. In prokaryotes (Bacteria, Archaea), the respiratory chain is located in the cytoplasmic membrane. The respiratory chain of the bacterium Paracoccus denitrificans appears to have many similarities to the mitochondrial type (Harold, 1986). However, many bacteria have variations compared to the mitochondrial type presented in Figure 1. Energetics and ATP conservation Most reducing equivalents are fed into the respiration process via NADH2, which has a standard redox potential (E00 ) of 0.32 V. The terminal electron-accepting redox pair O2/H2O has redox potential of þ0.82 V. Thus, the usable redox potential is about 1.14 V. From this redox potential one can calculate a standard DG00 of 440 kJ per mol of O2. Assuming that ATP conservation requires about 75 kJ per mol, the conservation of 5 to 6 ATP per O2 reduced is possible. As a consequence of the high ATP yield, aerobic organisms can form more biomass from the same amount of substrate than anaerobic organisms. Use of oxygen as cosubstrate Molecular oxygen is used as a cosubstrate in some catabolic and anabolic reactions (Lengeler et al., 1999). The enzymes involved are called mono- or di-oxygenases, depending on the number of oxygen atoms transferred to their substrate. Oxygenases are used by aerobes to activate certain substrates that do not have functional groups that allow a simple degradation. They catalyze the initial hydroxylation of hydrocarbons to yield alcohols that can easily be metabolized further. They are also involved in the ring cleavage of aromatic compounds and in the first oxidation step of ammonia oxydation. Anaerobic organisms are either unable to oxidize the substrates activated

AEROBIC METABOLISM

9

either the products are absent in strict anaerobes (e.g., sterols), or anaerobes have found alternative pathways for their biosynthesis.

Toxic forms of oxygen and detoxification Aerobic metabolism inevitably generates small amounts of partially reduced toxic oxygen species (Lengeler et al., 1999). These are the superoxide anion (O2.), hydrogen peroxide (H2O2), and the hydroxyl radical (.OH). These species can be formed via nonenzymatic one-electron reduction steps (e.g., with Fe2þ, quinols, or H2S) or by nonspecific reactions with reduced flavoproteins. Aerobic bacteria can remove toxic oxygen species by nonenzymatic reactions (e.g., with glutathione) or by means of enzymes: Superoxide dismutase (SOD) converts superoxide to H2O2, and catalyze dismutates H2O2 to water and O2. These enzymes often lack in strict anaerobes, which might explain their sensitivity toward oxygen. Distribution of the aerobic lifestyle Many microorganisms can thrive in oxic (oxygencontaining) as well as in anoxic (free of oxygen) environments. Those are classified as facultative aerobes (or facultative anaerobes). This means that they can rely on aerobic respiration as well as on respiration with other electron acceptors or on fermentative energy conservation. Groups of strict anaerobes comprise methanogens, sulfate reducers, and Clostridia. Even in strict anoxic (oxygen-free) environments facultative aerobes can be found (Süß et al., 2006).

Aerobic Metabolism, Figure 1 Principle of respiratory electron transport and vectorial proton translocation. The first proton translocating complex (with FMN and FeS as prosthetic groups) channels electrons to the quinones (UQ), which perform a cycle, again coupled to proton translocation. Electrons move on to the cytochrome bc1-complex, via cytochrome c to the coppercontaining terminal cytochrome oxidase (Cyt aa3). O2 is reduced in the last step only. The ATP synthesis coupled to reentry of the translocated protons is independent of the electron transport process. (From Cypionka, 2006.)

by means of oxygenases or possess different degradation pathways. Anabolic reactions making use of molecular oxygen are the biosynthesis pathways of tetrapyrroles, ubiquinones, pyrimidines, sterols, and unsaturated fatty acids. Again,

Summary Aerobic respiration is the most effective way of energy conservation. It is used to oxidize the major part of the organic matter on Earth. During aerobic respiration oxygen is reduced to water. In some cases, O2 is used by oxygenases for the hydroxylation of inert substrates. Aerobic organisms have enzymatic and nonenzymatic mechanisms to detoxify partially reduced oxygen species such as superoxide or hydrogen peroxide. Bibliography Cypionka, H., 2006. Grundlagen der Mikrobiologie. Heidelberg: Springer. Harold, F. M., 1986. The Vital Force: A Study of Bioenergetics. New York: Freeman and Company. Lengeler, J. W., Drews, G., and Schlegel, H. G., 1999. Biology of the Prokaryotes. Stuttgart: Thieme. Mitchell, P., 1966. Chemiosmotic coupling in oxidative and photosynthetic phosphorylation. Biological Reviews of the Cambridge Philosophical Society, 41, 445–502. Süß, J., Schubert, K., Sass, H., Cypionka, H., Overmann, J., and Engelen, B., 2006. Widespread distribution and high abundance of Rhizobium radiobacter within Mediterranean subsurface sediments. Environmental Microbiology, 8, 1753–1763.

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ALGAE (EUKARYOTIC)

Cross-references Anaerobic Transformation Processes, Microbiology Archaea Bacteria Carbon (Organic, Degradation) Critical Intervals in Earth History

ALGAE (EUKARYOTIC) Thomas Friedl1, Nicole Brinkmann1, Kathrin I. Mohr1,2 University of Göttingen, Göttingen, Germany 2 Helmholtz Centre for Infection Research, Braunschweig, Germany 1

Algae (eukaryotic) Eukaryotic algae are a collection of extremely diverse, nonrelated organisms that perform photosynthesis in plastids, permanent organelles of green, brown, or bluish colors derived from endosymbiosis. In contrast to plants, algae do not form embryos. Algae is a term of convenience and refers to a collection of highly diverse organisms that undertake photosynthesis and/or possess plastids (Keeling, 2004). Many authors even include the prokaryotic cyanobacteria into the algae, because they exhibit a life-style rather similar to their eukaryotic counterparts and often share the same habitat with eukaryotic algae. Cyanobacteria form the origin of plastids (for reviews see McFadden, 2001; Keeling, 2004; Palmer, 2003). Plastids are the organelles of plants and eukaryotic algae that harbor photosynthesis and synthesize many chemical compounds also important for other biochemical pathways (e.g., aromatic amino acids, heme, isoprenoids, and fatty acids); nonphotosynthetic plastids are known from plants and many algal groups (Williams and Keeling, 2003). Recently, from strategies of synthesizing various information, that is, mainly DNA sequence analyses of concatenated genes as well as the incorporation of discrete characters such as insertions, deletions, and gene fusion events, and consideration of morphology and biochemistry, a phylogeny of eukaryotes has emerged which consists of five “supergroups” (Keeling, 2004). Algae are scattered among four of the five major eukaryotic groups. The “Plantae” comprises exclusively eukaryotes with plastids derived from primary endosymbiosis, that is, they are bound by two membranes, derived from the inner and outer membranes of a cyanobacterium which was transformed into an organelle through uptake and retention by the host cell followed by the loss of much of its genome (Keeling, 2004). The Plantae are formed by three distinct but related lineages, the viridiplantae (which comprises all green algae and land plants or embryophytes), the rhodophytes (red algae), and the glaucophytes. The “Chromalveolates” comprises large and abundant algal groups such as the heterokont algae (e.g., diatoms and brown algae), the haptophytes,

the alveolates (which include the dinoflagellates and apicomplexans), and the cryptomonads. Only a single lineage of each of the “Excavates” and “Rhizaria” includes algae, the euglenoids and chlorarachniophytes. Except for the Plantae, all other eukaryotic algal lineages acquired their plastids through secondary endosymbiosis (for reviews see McFadden, 2001; Keeling, 2004; Palmer, 2003). Secondary plastids are surrounded by four or three membranes which resulted from phagocytosis of a primary alga, which was either a red alga or a green alga. In secondary plastids of both red and green origin, the primary algal nucleus is highly reduced (in the cryptomonads and chlorarachniophytes) or more commonly lost altogether (e.g., in the heterokonts and haptophytes; Keeling, 2004).

Green algae (Plantae supergroup) The green algae are photosynthetic eukaryotes with double membrane-bound plastids (derived from primary endosymbiosis), stacked thylakoids, and the chlorophylls a and b. Their other accessory pigments are the carotenoids, beta-carotene, and xanthophylls. The main reserve polysaccharide is starch which is deposited inside the plastid (Lewis and McCourt, 2004; Pröschold and Leliaert, 2007). A few genera (e.g., Prototheca) contain nonphotosynthetic plastids which lost their pigments secondarily. Apart from plastid-associated features, a unique stellate structure linking nine pairs of microtubules in the flagellar base of those green algae that are motile or produce motile stages is characteristic, as well as cell walls composed of cellulose which is present in most green algae (Graham et al., 2009). Most of these characters are shared with embryophytes (land plants) and, therefore, the green algae are difficult to define to the exclusion of the latter. The green algae have evolved in two major lineages (Friedl, 1997; Karol et al., 2001; Lewis and McCourt, 2004). One lineage, the chlorophyte lineage (Chlorophyta sensu Bremer, 1985; Sluiman, 1985) comprises the majority of green algal diversity (i.e., the green algae in a more narrow sense or the majority of what have been traditionally called green algae) with a bewildering array of morphologies attributable to at least five different morphological organizations (Pröschold and Leliaert, 2007). Examples of microscopic green algae reported from freshwaters with high calcification levels are rather small coccoid forms (e.g., Chlorella-like, Figure 1a; Desmochloris, Figure 1d; Pseudendocloniopsis, Figure 1f ), coccoids that form coenobia (e.g., Scenedesmus, Figure 1b) or are of filamentous organization which may easily disintegrate into unicells (e.g., Stichococcus, Figure 1c) or form small thalli of branched filaments (e.g., Pseudendoclonium, Figure 1e). Also some monadoid (flagellated unicells) green algae and the siphonocladous Cladophora are well known from highly calcifying freshwater habitats. Several marine green algae of coenocytic organization precipitate carbonate and belong to the

ALGAE (EUKARYOTIC)

a

b

c

d

e

f

g

h

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Algae (Eukaryotic), Figure 1 Eukaryotic algae recovered from tufa stromatolite biofilms of hardwater creeks: (a) Chlorella-like coccoid green alga (Trebouxiophyceae, pure culture), (b) coenobia of coccoid green alga Scenedesmus sp. (Chlorophyceae, pure culture), (c) barrel-shaped unicells of green alga Stichococcus sp. (Trebouxiophyceae, pure culture), (d) coccoid green alga Desmochloris sp. (Ulvophyceae, pure culture), (e) branched filaments of green alga Pseudendoclonium sp. (Ulvophyceae, pure culture), (f) coccoid green alga Pseudendocloniopsis sp. (Ulvophyceae, pure culture), (g) filamentous unbranched stramenopiles Xanthonema sp. (Xanthophyceae, pure culture), (h) filamentous branched Heterococcus sp. (Xanthophyceae, pure culture), scale bar 20 mm.

calcareous algae, for example, the orders Dasycladales (Kingsley et al., 2003) and Caulerpales (Stanley et al., 2010). Whether the aforementioned freshwater green algal taxa are involved in carbonate precipitation is still

unknown, but microscopic freshwater green algal genera with calcite deposits on their surfaces are several members of Zygnematales (e.g., Oocardium) and Gongrosira (Rott et al., 2009; Freytet and Verrecchia, 1998). The other major

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green algal lineage, the charophyte clade (sensu Lewis and McCourt, 2004), contains a smaller number of widespread, mostly freshwater green algae. Examples from the charophyte clade which are associated with calcification are the orders Zygnematales and Charales, for examples, the genus Chara is known for its extracellular carbonate production (McConnaughey, 1991). The charophyte green algae represent the closer relatives of land plants (embryophytes) with the Charales being most closely related to embryophytes (Karol et al., 2001; Lewis and McCourt, 2004). In the three lineages of green algae where calcite accumulation is known, that is, the orders Caulerpales, Dasycladales and Zygnematales (Bilan and Usov, 2001), aragonite deposits are outside of the cell wall but inside a specialized sheath. The crystal bundles are organized in layers, which are parallel to the filaments of this porous multilayer sheath and have also calcium oxalate inclusions. The chemical composition of the sheath and its role in the calcification process are unknown for the majority of species. It may simply be a barrier preventing the diffusion into the surrounding medium, although its direct participation in the initiation of the crystal formation cannot be excluded (Bilan and Usov, 2001). A third group of green algal taxa, called prasinophytes, consists of “primitive”-appearing unicells of uncertain affinity. Most known prasinophyte green algae form several early diverging clades within the phylogeny of the Chlorophyta (Marin and Melkonian, 2010). The green flagellate genus Mesostigma may represent a very basal lineage of the charophyte clade (Charophyta), but some prasinophytes are likely representatives of even other early diverging clades (Fawley et al., 2000). With respect to the systematics of green algae, there is now consensus that the majority of known green algal species of the Chlorophyta belongs to three major lineages, the chlorophytes, trebouxiophytes, and ulvophytes, which are assigned to class rank (classes Chlorophyceae, Trebouxiophyceae, and Ulvophyceae; Pröschold and Leliaert, 2007). Green algae have an affinity to form symbioses with hosts of various phylogenetic positions. Well-known examples are green algae forming the photoautotrophic partners (photobionts) of fungal symbioses, that is, lichens, or representing the algal hosts (“Zoochlorellae”) of ciliates (Alveolates), heliozoa, sponges, and sea anemones (e.g., Summerer et al., 2008; Letsch et al., 2009).

Rhodophytes – red algae (Plantae supergroup) Red algae are a very large and diverse group of microscopic algae and macroalgae. They are best known for their economic and ecological importance. Though they are present in freshwater with several genera, the majority of red algal genera occur on tropical and temperate marine shores, where they play important ecological roles (Graham et al., 2009). Certain calcified red algae, known as corallines, form hard, flat sheets

that consolidate and stabilize coral reef crests, that is, coralline red algae protect reefs from wave damage and, thus, they are regarded as important keystone organisms. Of well-known economic importance are the species of Porphyra and other red algal species which are grown in mariculture operations for use as human food. Several other marine genera are cultivated or harvested for the extraction of gelling polysaccharides such as agarose and carrageen (Graham et al., 2009). Corallines are known from fossil records 500 million years old (Lower Ordovician; Riding et al., 1998). The oldest fossil record of a (non-coralline) red alga may represent the fossil taxon Bangiomorpha pubescens which was described from the 1,200-million-year-old Hunting Formation in the Canadian Arctic (Butterfield, 2000). It is most notable that B. pubescens even represents the earliest putative record for sex and taxonomically resolvable complex multicellularity among eukaryotes (Saunders and Hammersand, 2004). For a review of fossil records of the various taxonomic groups of red algae, see Saunders and Hammersand (2004). The plastids of red algae lack chlorophyll b and c, but contain phycobilins (allophycocyanin, phycocyanin, and the red pigment phycoerythrin) as accessory pigments which are located in phycobilisomes on the outer surface of the thylakoids. The red algal plastids are bounded by two membranes and derived from a cyanobacterial primary endosymbiosis (Keeling, 2004). They produce floridean starch which is deposited in the cytoplasm. The rhodophytes lack flagella and centrioles in all stages of their life history (Graham et al., 2009). The red algae (Rhodophyta) are a distinct eukaryotic lineage, whose members are united in the phylogenetic analyses of a nuclear, plastid, and mitochondrial genes (for review see Yoon et al., 2006). They belong to the Plantae supergroup of eukaryotes (Keeling, 2004) and are believed to form a sister group with the green line (Martin et al., 2002). Their divergence has been estimated ca. 1,500 million years ago (Yoon et al., 2004). Based on molecular evidence, the red algae are partitioned into seven classes, six of which have a rather simple body structure and reproduction (Yoon et al., 2006). The florideophytes (class Florideophyceae) are the monophyletic group of red algae that exhibit more complex bodies and reproduction (Saunders and Hammersand, 2004). The ecologically significant corallines, the economically important genus Chondrus (used as food and for the extraction of carrageen; Graham et al., 2009), and the vast majority of other modern red algal species are florideophytes; they are divided into several monophyletic groups (subclasses; Saunders and Hammersand, 2004; Le Gall and Saunders, 2007). Coralline red algae represent not only a large carbon reservoir, but they also consolidate sediment and build landforms, and they occupy harder substrate in the

ALGAE (EUKARYOTIC)

world’s oceans than any other group of photosynthetic organisms. Unlike most other calcified algae that deposit carbonate as aragonite, the coralline red algae form calcite and it is believed that each family of coralline red algae is defined by only one crystalline modification of calcium carbonate (Borowitzka, 1977). The red algae show evidence of a surprising variety of calcification mechanisms, but the location of deposition is even more diverse. Mineralization may occur throughout the walls of nearly all vegetative cells, it may occur within the walls of only specialized cells, or it may be located in an intercellular organic matrix between cells, but not within the cell walls (Pueschel et al., 1992). Calcification in corallines is poorly understood, however, it seems clear that corallines deposit high magnesium calcite in a definitive pattern within their cell walls via the membrane pumps that control the flux of calcium and carbonate with the control of crystal orientation by organic wall components (Zankl, 2007). Coralline red algae can be the deepest algae (50 to 110 m, Dullo et al., 1990) and many species occur in great abundance into the Arctic and subarctic. Coralline red algae in coral reefs are ubiquitous and dominant (Glynn et al., 1996; Keats et al., 1997), and their abundance in cryptic and shaded environments can be greatly underestimated (Littler, 1972). In temperate systems, the accumulation of unattached living or dead coralline algae forms large maerl beds (Martin et al., 2006). They also provide food for herbivores with hardened mouthparts (Steneck and Dethier, 1994) and surfaces for the settlement of invertebrate larvae (Adey, 1998). Coralline algal abundance, size, shape, and species composition in maerl beds vary considerably depending on their location, and several environmental factors influence coralline algal distribution. Maerl beds are highly sensitive to desiccation and are found from the low intertidal zone to depths of 150 m (Foster, 2001). Furthermore, coralline species are sensitive to seasonal variations in temperature inducing changes in their physiology, for example, Lithothamnion spp. growth rate increases with water temperature (Potin et al., 1990). Despite their ecological importance in temperate systems, primary production and calcification of coralline algae has been mainly investigated in tropical areas (Chisholm, 2003; Payri et al., 2001). A number of different associations between sponges and several species of red macroalgae have been described in literature (Trautman et al., 2000, 2002). The symbiotic association between the haplosclerid sponge Haliclona cymiformis and the red macroalga Ceratodictyon spongiosum is common on shallow coral reefs of the Indo-West Pacific, where it has been described as being one of the most conspicuous organisms found in these areas (van Soest, 1990). This symbiosis seems to be an obligatory association as neither sponge nor alga has ever been identified growing independently or in association with other species (Morrissey, 1980).

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Glaucophytes (Plantae supergroup) Glaucophytes (or glaucocystophytes) form a small group of microscopic algae exclusively found in freshwater environments; only about 13 species of glaucophytes have been described so far. They represent the third lineage of the plant supergroup, which is defined by plastids bound by a double membrane, that is, which is derived from primary cyanobacterial endosymbiosis (Keeling, 2004). Glaucophyte plastids are unique among plastids in that they are retaining the prokaryotic peptidoglycan layer between their two membranes. Therefore, they may represent an intermediate in the transition from a cyanobacterial endosymbiont to plastid; molecular phylogenetic analyses show the glaucophytes as the earliest divergence within the plant supergroup (Keeling, 2004; Martin et al., 2002). Glaucophyte plastids share the accessory photosynthetic pigments, phycobilins, which are organized in phycobilisomes (small particles on the outer surface of thylakoid membranes) with cyanobacteria and rhodophytes. For a review on glaucophytes, see Bhattacharya and Schmidt (1997) and Steiner and Löffelhardt (2002). Photosynthetic stramenopiles (chromalveolates) Stramenopiles form a monophyletic group of photoautotrophic as well as heterotrophic organisms which are characterized by swimming cells possessing at least one flagellum with distinct tripartite tubular hairs (Graham et al., 2009; Andersen, 2004). Most stramenopiles have two distinctly different flagella and, therefore, are also known as heterokonts. They have a long flagellum with tripartite hairs (tinsel or immature flagellum) to pull the cells through the water and a shorter smooth one (whiplash or mature flagellum) that lacks tripartite hairs, often with a light-sensing flagellar swelling at its base (Graham et al., 2009; Andersen, 2004). Photosynthetic stramenopiles or heterokont algae usually appear brown or golden brown (and are thus sometimes referred to as chromophytic algae) due to the presence of characteristic accessory pigments, fucoxanthin or vaucheriaxanthin, and at least one form of chlorophyll c (except in eustigmatophytes). The plastids are derived from a secondary endosymbiosis event with a red alga involved and surrounded by four membranes (McFadden, 2001; Keeling, 2004; Palmer, 2003). In most photosynthetic stramenopiles, a chloroplast endoplasmatic reticulum is present, that is, plastid and nucleus are structurally connected. The photosynthetic stramenopiles comprise a tremendous morphological diversity: a large variety of unicellular or colonial algae, including the diatoms with silica frustules as walls, multicellular simple filaments (e.g., in the xanthophytes) as well as the macroscopic complex brown algae forming a brown canopy at sea shores. Based on detailed analyses of pigment composition, ultrastructure and molecular phylogenies more than a dozen of different classes of photosynthetic stramenopiles have been described (Graham

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et al., 2009; Andersen, 2004). Some photosynthetic stramenopiles do not form mineralized cysts or scales and, therefore, they have no fossil record. Examples are the brown seaweeds (Phaeophyceae), the yellow green algae (Xanthophyceae), and the inconspicuous eustigmatophytes. In the context of geobiology also xanthophytes and eustigmatophytes may be important, as they have been reported from freshwaters with high calcite precipitation (see Figure 1g, e; Arp et al., 2010). However, their role in the calcification processes is still unclear. Therefore, in this entry, only those classes of photosynthetic stramenopiles will be discussed that produce mineralized cell coverings which are important in fossil deposits and, therefore, may be of particular interest in geobiology. Siliceous frustules define the diatoms (Bacillariophyta; see entry “Diatoms”), silica skeletons are characteristic for silicoflagellates (class Dictyochophyceae) and silica scales and cysts are commonly formed in synurophytes and chrysophytes. Silicoflagellates are unicellular, flagellated, and photoautotrophic stramenopiles which are characterized in at least one life cycle stage by the formation of distinctive, onepiece external silica skeleton that is highly perforate. They form the order Dictyochales, one of three orders that have been described for the class Dictyochophyceae (Daugbjerg and Henriksen, 2001; Graham et al., 2009). Siliceous skeletons from silicoflagellates are common in fossil deposits dating back to the middle Cretaceous (ca. 120 million years ago). The skeletal remains in fossil material also indicate that species diversity reached a peak in the Miocene (ca. 23–25 million years ago), with more than 100 described species, but the diversity of silicoflagellates has decreased thereafter, leaving behind only two genera (Daugbjerg and Henriksen, 2001; Tappan, 1980). Biostratigraphic zonations have been developed for silicoflagellates in all oceans throughout the Cenozoic, though the zones typically range over longer intervals of time than foraminiferal or diatom zones (McCartney and Wise, 1990; McCartney and Harwood, 1992). Silicoflagellate skeletons may have considerable potential as ecophenotypic indicators (McCartney and Wise, 1990). Modern silicoflagellates are widespread in the oceans with a tendency to be more abundant in colder waters where they can even grow up to form blooms (Graham et al., 2009). Dictyochophytes occur in both marine and freshwater habitats (Moestrup, 1995; Moestrup and O’Kelly, 2000). Synurophytes and chrysophytes (often summarized as “Chrysophyte Algae”) produced an exceptional fossil record owing to the high preservation potential of their siliceous stomatocysts and scales (Smol, 1995; Andersen, 2004; Graham et al., 2009). In addition, they are highly sensitive to an array of limnological parameters. Synurophytes and Chrysophytes are close relatives in molecular phylogenies (Andersen, 2004); both share the capability to form stomatocysts, silica-walled resting stages, which arise sexually or asexually under unfavorable environmental conditions. The walls of stomatocysts

are so heavily silicified that they resist silica dissolution processes and thus accumulate in the sediments of lakes (Graham et al., 2009). Fossil records of stomatocysts extend back to the Lower Cretaceous (at least 150 million years ago). Synurophytes and chrysophytes are able to switch between autotrophy, heterotrophy, and even phagotrophy, which puts them at a distinct advantage in aquatic ecosystems that are low in nutrient concentrations and have reduced light penetration (Betts-Piper et al., 2004). From patterns of stomatocyst assemblages, past environmental conditions can be inferred making stomatocysts very important for paleolimnology (Duff et al., 1995; Siver, 1995; Wilkinson et al., 2001; Zeeb and Smol, 2001). Synurophytes are unicellular or colonial silicated flagellates and despite they are photoautotrophic, many species are also capable of phagotrophy. Synurophycean cell surfaces are covered by overlapping silica scales, sometimes with spiny bristles, and are perforated (Graham et al., 2009). The scales’ perforation patterns are used as taxonomic features to delimitate species of synurophytes. A number of adhesive polysaccharides are involved in the scale attachment. The scales are produced in certain vesicles located on the surface of one of the plastids. In contrast to diatoms, synurophytes can continue to divide and function also in the absence of an external silica covering. In cultures of naked cells most cells will recover a complete cell covering when silicate is resupplied to silica-depleted cells (Leadbeater and Barker, 1995). Fossil scales similar to those of modern synurophytes have been reported from deposits of the middle Eocene (about 47 million years ago; Siver and Wolfe, 2005). Synurophyte scales can be used like stomatocysts for tracking environmental change using lake sediments (Zeeb and Smol, 2001; Wolfe and Perren, 2001). Synurophytes are abundant in neutral to slightly acidic freshwaters. Some species are regarded as indicators of low levels of pollution, but others are characteristic of eutrophic lakes (Graham et al., 2009). Chrysophytes are golden-brown microalgae that are mostly unicellular or colonial which occur as flagellates or nonmotile cells; they include many unique and interesting morphologies (Graham et al., 2009). In contrast to synurophytes, scales covering the cell surface are absent in chrysophytes. Chrysophytes typically favor slightly acidic freshwaters of moderate to low productivity; their abundance and species richness increase with lake eutrophic status (Elloranta, 1995). Chrysophytes may have strong ecological impacts because many are mixotrophs, able to take up and metabolize dissolved organic compounds and particulate food as well as photosynthesize (Graham et al., 2009). Several chrysophytes are associated with the formation of undesirable blooms because living cells of certain species can produce toxic fatty acids that affect fish or excrete aldehydes and ketones into the water which can give it an unpleasant taste and odor.

ALGAE (EUKARYOTIC)

Coccolithophorids and haptophyte algae (chromalveolates) Haptophyte algae are a monophyletic group that includes all photosynthetic organisms with a haptonema. However, haptophytes also include some nonphotosynthetic relatives, and some that have secondarily lost the haptonema (Andersen, 2004). The haptonema, from which the group derives its name, is a microtubule-supported appendage that lies between two approximately equal flagella (for a review, see Inouye and Kawachi, 1994). Most haptophytes are marine, occurring in significant abundances even at greater water depth (up to 200 m). Haptophyte algae share many features, for example, pigment composition, presence of a chloroplast endoplasmatic reticulum, and plastids derived from red algal secondary endosymbiosis with the photosynthetic stramenopiles (Andersen, 2004). Based on structural and molecular evidences the haptophytes form a monophyletic lineage, treated as phylum Haptophyta which is divided into two classes (Edvardsen et al., 2000; Andersen, 2004). The larger group, class Coccolithophyceae, comprises the coccolithophorids which are haptophytes with a cell coat of mineralized scales (coccoliths) and two equal flagella or lack flagella altogether. Coccoliths are largely composed of calcium carbonate crystals in the form of calcite. There are two types of coccoliths, holococcoliths and heterococcoliths. In case of holococcoliths, an organic scale consisting largely of cellulose and produced by the Golgi body is secreted to the cell surface and then calcite crystals which are held together by organic material are deposited on the scale. In contrast, heterococcoliths develop on organic scales before being secreted from the cell, that is, calcite crystal deposition is already within the cell. The calcite crystals grow and interlock gradually forming the complex shape of the mature coccolith (deVrind-deJong et al., 1994; Probert et al., 2007; Graham et al., 2009). Coccoliths do not degrade at normal ocean pH and thus accumulate on the ocean floor. Therefore, coccolithophorids have produced huge amounts of sedimentary carbonates. Coccolithophorids remove large quantities of atmospheric CO2 through their photosynthesis and calcification and, therefore, are an important component of the global carbon cycle (McConnaughey, 1994), accounting for a substantial part of the ocean floor limestone sediments. Coccolithophorids contribute at least 25% of the total annual vertical transport of inorganic carbon to the deep ocean (Rost and Riebesell, 2004). Coccolithophorids have an excellent fossil record with fossil coccoliths well over 200 million years old. The abundance of coccolith fossils peaked during the late Cretaceous (63–95 million years ago) when extensive chalk deposits were laid down. Coccoliths are widely used as stratigrafic indicators and fossil coccoliths are widely used as bioindicators in the oil industry and as indexes of past climate and ocean chemistry conditions (Young et al., 1994).

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Haptophytes are ecologically significant in terms of both biotic interactions and biogeochemistry. Haptophyte algae are considered as high-quality foods for zooplankton; several species contain nutritionally important polyunsaturated fatty acids which make them also commercially valuable for the production of fish in aquaculture systems. Other coccolithopytes may produce toxins that destroy cell membranes or produce copious amounts of organic slime and foam (Graham et al., 2009). Many coccolithophorids form blooms in ocean waters and produce large amounts of a volatile sulfur-containing molecule, dimethyl sulfide (DMS) that enhances cloud formation and increases acid rain (Malin and Steinke, 2004). Another cooling effect on the climate comes from the coccoliths which readily reflect light, thereby increasing reflectance of the ocean’s surface.

Dinoflagellates (chromalveolates) The dinoflagellates are an important group of phytoplankton in marine and freshwaters. Their adaptation to a wide variety of environments is reflected by a tremendous diversity in form and nutrition and an extensive fossil record dating back several hundred million years (Graham et al., 2009). Dinoflagellate cells have two structurally distinct flagella whose motion causes the cells to rotate as they swim, but also nonflagellate unicells and filamentous forms of dinophytes are known. Two different cell types can be distinguished on the basis of the cell-wall covering or theca. The “naked” or unarmored forms have an outer plasmalemma surrounding a single layer of flattened vesicles (membrane sacs or alveoli). Armored dinoflagellates have cellulose or other polysaccharides within each vesicle, giving the cells a more rigid, inflexible wall. These cellulose plates are arranged in distinct patterns which are extensively used as taxonomic “fingerprints” (Hackett et al., 2004). Cells of armored dinoflagellates display conspicuous anterior–posterior and dorsal–ventral differentiation. Their cells consist of two parts, separated by a groove that encircles the cell and contains the transverse flagellum which is flattened, ribbon-shaped, and with a single row of hairs. A smaller groove is mostly extending into the posterior cell part and contains the longitudinal flagellum which has two rows of hairs; it is directed posterior and emerges from the cell (Graham et al., 2009; Hackett et al., 2004). Success of dinoflagellates as phytoplankton may be due in part to unique behavior patterns, including duel vertical migration and their capability to live phagotrophic (feeding on particles such as the cells of other organism) in addition to photoautotrophy (mixotrophic life-style). Some of the armored or thecate heterotrophic dinoflagellates have developed a remarkable pseudopod-like structure that is extruded from the cell and flows around the prey, enveloping it so the contents can then be digested (Hackett et al., 2004). Some dinoflagellates produce toxins that are dangerous to man, marine mammals, fish,

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seabirds, and other components of the marine food chain (Van Dolah, 2000). Others are bioluminescent and emit light; some function as parasites or symbionts that rely on host organisms for part of their nutrition. Many photosynthetic dinoflagellates live as endosymbionts with reefbuilding corals. Stimulated by chemical signals from their animal hosts, dinoflagellate endosymbionts provide the corals with essential organic food. Therefore, the dinoflagellate symbionts are critical components of the coral reef ecosystems whose loss during stress-related “bleaching” events can lead to mass mortality of coral hosts and associated collapse of reef ecosystems (Baker, 2003). Also, the high rates of calcification and production that characterize typically oligotrophic coral reef ecosystems are largely credited to the mutualistic symbiosis between the scleractinian corals and photosynthetic dinoflagellates belonging to the genus Symbiodinium (Apprill and Gates, 2007). Corals with a dinoflagellate symbiont calcify much faster than those without – an effect linked to photosynthetic fixation of CO2 by the dinoflagellates (Marshall, 1996). A significant amount of photosynthetic product is excreted by the symbiotic dinoflagellates, primarily as glycerol. Up to 50% of the fixed carbon may be transferred to the host (Paracer and Ahmadjian, 2000), in which it is converted mainly to lipids and proteins. On the dinoflagellate side, many of these symbioses occur in oligotrophic waters in which nutrients are scarce in the water column (Hackett et al., 2004). In other symbiotic dinoflagellate associations, the hosts include foraminifera, radiolarians, flatworms, anemones, jellyfish, and even bivalve mollusks (Hackett et al., 2004). Undisputed structural fossils of dinoflagellates occur some 200 million years or so ago (Graham et al., 2009). These fossils, known as hystrichospheres, resemble resting stages of some modern dinoflagellates. Some modern dinoflagellates produce resting cysts or zygotes with an organic compound resistant to decay and thus aid the survival of dinoflagellate fossils, of which a considerable diversity has been described (Fensome et al., 2003). Other chemical compounds thought to be specific to dinoflagellates have already been found in Proterozoic (Precambrian) rocks and such chemical fossils suggest that dinoflagellate-like organisms might have existed already more than 600 million years ago (Graham et al., 2009). Many dinoflagellates are photosynthetic and, through endosymbiosis, have acquired a wide diversity of plastids from distant evolutionary lineages. The most common plastid in dinoflagellates is golden brown with a unique accessory pigment, peridinin; they also contain chlorophyll c. Such peridinin plastids are bounded by three membranes and are derived from red algal secondary endosymbiosis (Delwiche, 2007; Hackett et al., 2004). A number of dinoflagellates have plastids from haptophytes, diatoms, green algae, or cryptomonads which may have replaced preexisting peridinin plastids (tertiary endosymbiosis; McFadden, 2001; Keeling, 2004; Palmer, 2003).

A small group of dinoflagellates with peridinin plastids produce calcareous structures, formed during the life cycle or found in vegetative stages (Gottschling et al., 2005). The potential to produce calcareous structures has been considered as apomorphic within alveolates (Kohring et al., 2005), arguing for the monophyly of the family Calciodinellaceae which comprises approximately 30 (recognized) extant species that are distributed in cold through tropical seas of the world. Calcareous cysts are deposited in both sediments coastal (Montresor et al., 1998; Persson et al., 2000) and oceanic (Vink, 2004; Zonneveld et al., 1999). According to the fossil record, calcareous dinoflagellates originate in the Upper Triassic (Janofske, 1992) and are highly diverse during the Cretaceous and throughout the Tertiary (Keupp, 1991; Kohring, 1993; Willems, 1994), thus many fossil species (namely their cysts) have been described (Gottschling et al., 2005). Based on morphological and molecular data, calcareous dinoflagellates (Thoracosphaeraceae, Peridiniales) are a monophyletic group comprising three major clades (Gottschling et al., 2008).

Other chromalveolates Cryptomonads. Recent molecular phylogenetic analyses indicate that the cryptomonads and the haptophyte algae together form a monophyletic group (Hackett et al., 2007) and, therefore, also belong to the Chromalveolates supergroup. Cryptomonads form an abundant group of marine and freshwater unicellular flagellates that contain a plastid derived from red algal secondary endosymbiosis with chlorophyll a and c surrounded by four membranes. Cryptomonad plastids have received some attention because they are one of only two groups in which the primary algal nucleus has not been completely lost; they retain a small relict nucleus called a nucleomorph (McFadden, 2001; Keeling, 2004; Palmer, 2003). Apicomplexa are a large group composed entirely of obligate intracellular parasites, including several that cause significant diseases such as malaria. As intracellular parasites, the discovery of a relict plastid (or apicoplast) in apicomplexa bounded by four membranes has drawn a great deal of attention as an evolutionary novelty and possible drug target (Keeling, 2004). There is current evidence indicating that this plastid is derived from a red alga and that the apicomplexa share a monophyletic origin with all other groups with plastids from red algal secondary endosymbiosis. The apicomplexa may be most closely related with the dinoflagellates (Moore, 2008). For a review of apicomplexa relating to their plastids, see Foth and McFadden (2003). Algal groups with plastids derived from green algal secondary endosymbiosis Chlorarachniophytes (Rhizaria supergroup). Chlorarachniophytes are a relatively rare group of marine amoeboflagellates and flagellates that contain a green algal plastid (chlorophyll a and b) bounded by four membranes. They

ALGAE (EUKARYOTIC)

contain a relict nucleus, nucleomorph, which has been retained from the green algal symbiont (McFadden, 2001; Keeling, 2004; Palmer, 2003). Chlorarachniophytes form a monophyletic group within the Rhizaria supergroup of eukaryotes (Ishida et al., 1999). Euglenids (Excavates supergroup). Euglenids are a diverse group of common marine and freshwater flagellates, about half of which contain a plastid derived from green algal secondary endosymbiosis which is bounded by three membranes and contains chlorophyll b as accessory pigment. The remainder of the group are osmotrophs or heterotrophs that feed on bacteria or other eukaryotes. Molecular phylogenies show that photosynthetic euglenids may have acquired their plastids from a green alga relatively late in evolution, despite the Euglenozoa may be a rather old group. Euglenids are closely related to the parasitic trypanosomes (Kinetoplasids), together with diplonemids, making up the monophyletic Euglenozoa. Distinguishing features of euglenids are that they produce a storage carbohydrate which is a ß-1,3linked glucan (paramylon) in their cytoplasm and that they display a unique surface structure composed of parallel ribbon-like proteinaceous strips. For reviews of euglenids, see Leedale and Vickerman (2000), Milanowski et al. (2006) or Ciugulea and Triemer (2010).

Summary Algae is a term of convenience and refers to a collection of highly diverse organisms that undertake photosynthesis and/or possess plastids. In addition, the photoautotrophic cyanobacteria are regarded as algae by some authors. In a recent phylogeny of the eukaryotes, the algae are scattered among four of the five recognized supergroups. A single primary endosymbiosis with a cyanobacterium may have been the start of all eukaryotic algae which first diverged into the green algae, red algae, and glaucophytes (Plantae supergroup). The plastids of all other algal lineages derived from secondary endosymbiosis involving either a red algal cell (algae of the Chromalveolates supergroup) or green algal cells (euglenids and chlorarchniophytes, Excavates and Rhizaria supergroups). Many algae form symbioses with ciliates and some metazoa. Algae are very important primary producers in almost every aquatic or terrestrial habitats and are important key players within the carbon and silica cycles for the major part of Earth history. Many lineages have produced significant fossil records due to their ability to precipitate calcium carbonate on their cell surfaces (red algae, some dinoflagellates) or form calcified (haptophytes) or silicified body scales, silica endoskeletons, resting stages, or cell walls (photoautotrophic stramenopiles, including dictyochophytes, synurophytes, chrysophytes, and diatoms). Due to their high sensitivity to certain environmental parameters and their high preservation potential, these algae also play important roles as sedimentary bioindicators in tracking environmental changes.

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Zankl, H., 2007. The origin of High-Mg-Calcite microbialites in cryptic habitats of Caribbean coral reefs – their dependence on light and turbulence. Facies, 29, 55–60. Zeeb, B. A., and Smol, J. P., 2001. Chrysophyte scales and cysts. In John, P., Smol, H., Birks, J. B., and Last, W. M. (eds.), Tracking Environmental Change Using Lake Sediments. Vol 3. Terrestrial, Algal, and Siliceous Indicators. Dordrecht: Kluwer, pp. 203–223. Zonneveld, K. A. F., Höll, C., Janofske, D., Karwath, B., Kerntopf, B., Rühlemann, C., Willems, H., 1999. Calcareous dinoflagellate cysts as paleo-environmental tools. In Fischer, G., and Wefer, G. (eds.), Use of Proxies in Paleoceanography. Berlin: Springer, pp. 145–164.

Cross-references Cyanobacteria Diatoms Early Precambrian Eukaryotes Fungi and Lichens Microbialites, Modern Microbialites, Stromatolites, and Thrombolites Photosynthesis Protozoa (Heterotroph, Eukaryotic) Symbiosis

ALKALINITY Andreas Reimer, Gernot Arp University of Göttingen, Göttingen, Germany The term “Alkalinity,” commonly denoted by TA (also AT, ALK, and others) and then called titration alkalinity or total alkalinity, refers to a very important chemical concept in aquatic chemistry. Total alkalinity is one of the few measurable quantities in natural waters that allows, together with other properties, to calculate concentrations of single species of the carbonate system such as CO2, HCO3, CO32, H+, and OH (Wolf-Gladrow et al., 2007), and further on, the saturation state of the waters with respect to certain carbonate minerals. Therefore, alkalinity represents a hydrogeochemical key parameter for the understanding of carbonate precipitation, either biologically mediated or not, and its variation through the geological history.

Definition From its determination by titrimetric techniques, alkalinity has historically been defined as the number of equivalents of a strong acid required to neutralize 1 L of water at 20 C. The obtained end-point, corresponding to the second equivalence point of the carbonate system, was operationally defined and results were then commonly given as eq L1. The recent definition of alkalinity is based on the implementation of the Brønsted–Lowry concept of acids and bases by Rakestrow (1949) who defined alkalinity as the excess of bases (proton acceptors) over acids (proton donors). Considering seawater, alkalinity is mostly determined by the ions bicarbonate and carbonate, also by the borate ion, and to a smaller extent by OH– and H+ from

dissociation of water. Alkalinity then refers to the proton condition with reference to a zero level of protons defined by the species H2CO3, B(OH)3, and H2O, where the proton acceptors are HCO3, CO32, B(OH)4, and OH and the proton donor is H+. That is, alkalinity is the equivalent sum of bases that have one or two protons less than the reference species minus H+ (or H3O+) that has one proton more than H2O (Dickson, 1981; Stumm and Morgan, 1996; Zeebe and Wolf-Gladrow, 2001). The expression for the total alkalinity is thus presented as TA ¼ ½HCO3   þ 2½CO3 2  þ ½BðOHÞ4   þ ½OH   ½Hþ ;

where conceptually the right hand term can be divided up in carbonate alkalinity ðCA ¼ ½HCO3   þ 2½CO3 2 Þ as the most dominant contributor, borate alkalinity, and water alkalinity. Natural waters may contain various acid–base systems that can accept and donate protons other than or in addition to carbonic acid or boric acid. These complications have been taken into account currently in the most precise definition of total alkalinity of Dickson (1981) (compare also Dickson, 1992; DOE, 1994; and recent extension by Wolf-Gladrow et al., 2007). The definition is unequivocally based on a chemical model of occurring acid–base processes and to be independent of temperature and pressure expressed in gravimetric units (mol kg1). In Dickson’s concept, the total alkalinity (TA) is defined as the number of moles of hydrogen ion equivalent to the excess of proton acceptors over proton donors with respect to the proton condition defined by the value pK = 4.5 in 1 kg of sample. Bases formed from weak acids with a dissociation constant K  10–4.5 are considered proton acceptors, acids with K > 10–4.5 are proton donors, and the chemical species defining the zero level of protons of the various acid–base systems are H2CO3, B(OH)3, H2O, H2PO4, H4SiO4, NH4+, H2S, SO42, F, and NO2. From this definition the expression for the total alkalinity is presented as TA ¼ ½HCO3   þ 2½CO3 2  þ ½BðOHÞ4   þ ½OH  þ ½HPO4 2  þ 2½PO4 3  þ ½H3 SiO4   þ ½NH3  þ ½HS  þ . . .  ½Hþ   ½HSO4    ½HF  ½H3 PO4   ½HNO2  . . . where the ellipses stand for additional acid–base species (Wolf-Gladrow et al., 2007). An alternative definition for alkalinity derives when considering the charge balance of natural waters (Stumm and Morgan, 1996). From the principle of electroneutrality total alkalinity can be expressed in terms of the charge imbalance between the equivalent sum of conservative cations (base cations of strong bases) and the sum of conservative anions (conjugate bases of strong acids). For the total alkalinity of sea water dominated by

ALKALINITY

HCO3, CO32, and B(OH)4, the expression may be presented by TA ¼ ½Naþ  þ ½Kþ  þ 2½Mg2þ  þ 2½Ca2þ   ½Cl   ½Br   ½NO3    2½SO4 2 ; which is equivalent to the above definition in terms of proton acceptors and donors TA ¼ ½HCO3   þ 2½CO3 2  þ ½BðOHÞ4   þ ½OH   ½Hþ : In the charge balance expression quantities are defined to be conservative with respect to mixing and changes in temperature (T ) and pressure (P) provided that concentrations are on the gravimetric unit scale (mol kg1). Thus, from the TA expression entirely in terms of conservative ions TA is a conservative quantity too. In contrast, the concentrations of individual species of, e.g., the carbonate system may change with temperature and pressure because of the T–P dependency of the equilibrium constants. Following the charge balance approach, Zeebe and Wolf-Gladrow (2001) and Wolf-Gladrow et al. (2007) have derived an expression in terms of total concentrations of conservative ions and conservative total concentrations of the various acid–base species under concern in the definition of Dickson (1981) in terms of an acid–base balance. This derived expression for TA is denoted as the explicit conservative alkalinity expression (TAec). TAec ¼ ½Naþ  þ ½Kþ  þ 2½Mg2þ  þ 2½Ca2þ  þ . . .  ½Cl   ½Br   ½NO3    . . .  2TSO4  THF  TPO4  THNO2 þ TNH3 ; where total concentrations terms represent concentrations sums of the species of the acid–base systems TSO4 = [SO42] + [HSO4], THF = [F] + [HF], TPO4 = [H3PO4] + [H2PO4] + [HPO42] + [PO43], THNO2 = [NO2] + [HNO2], TNH3 = [NH3] + [NH4+], and ellipses stand for additional ions with minor concentrations. The definition of TAec has the advantage that each term in the expression is conservative with respect to T–P changes, even though single species in the total concentration terms may be not. In particular, the approach is very convenient to derive changes of TA due to certain physicochemical and biogeochemical processes such as nutrient uptake in biological production and release during remineralization.

Sources, changes, and ranges The primary source of alkalinity in surficial (and subsurface) waters is carbonic acid-controlled chemical weathering of carbonates, aluminum-silicate minerals like feldspars or Mg-Fe-silicates. The overall concentration and subsequent changes in alkalinity of water are therefore determined by the geology of the watershed, the

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predominant weathering reactions and ion exchange in the soils, and concomitant biogeochemical processes. TA as a conservative quantity will first alter with changes in salinity as a result of precipitation, evaporation, and mixing. Contrariwise to its increase from dissolution of limestone carbonates TA decreases when biogenic or nonbiogenic CaCO3 or other carbonate minerals precipitate. Further important changes are due to various biogeochemical processes. It has to be noted that total alkalinity does not change as a result of CO2 exchange with the atmosphere. Invasion or degassing of CO2 affects the adjustment of the carbonate species (ratio of [HCO3] to [CO32]) and the pH but not TA. Therefore, it is also not the consumption of CO2 during photosynthesis or release during aerobic respiration that brings about a change in TA, but the utilization or remineralization of nitrate and phosphate involved in the processes. Nitrogen assimilation in form of ammonia decreases TA and no change of TA results from molecular nitrogen uptake (Stumm and Morgan, 1996; Wolf-Gladrow et al., 2007). Several microbially mediated processes lead to a change in alkalinity where deviation in the charge balance is compensated by yield or consumption of H+ or OH. When oxygen is consumed in processes like nitrification, oxygenation of soluble ferrous iron to ferric oxide or sulfide oxidation from HS or pyrite total alkalinity decreases. Under anoxic conditions, remineralization of organic matter by denitrification, reduction of iron and manganese oxides and sulfate reduction frequently result in large increases of total alkalinity (Lerman and Stumm, 1989; Stumm and Morgan, 1996). Depending on the rock mineralogy of the catchment area and prevalent vegetation total alkalinity in streams and inland lakes commonly ranges below 3 mmol kg1. Very low alkalinity is bound to tributaries and lakes in crystalline rock areas or to acidic bog and peatland waters. Here, H+ released by biomass exceeding H+ consumption by soil weathering results in an excess of protons that is referred to as “mineral acidity” or in turn “negative alkalinity” (Stumm and Morgan, 1996). On the other hand, in hard-water creeks that gain their alkalinity preferably from the dissolution of limestone rocks TA may reach values of 4 mmol kg1 to 6 mmol kg1 close to their spring sites. In the oceans, TA typically ranges from 2.2 mmol kg1 to 2.5 mmol kg1 where changes with depth are mainly associated with the production and dissolution of calcareous organisms (Broecker and Peng, 1982). Higher TA is found in today’s only anaerobic ocean basin, the Black Sea, where alkalinity increases to about 4.5 mmol kg1 in the anoxic bottom waters in the course of oxidation of organic material by sulfate reduction (Kempe, 1990; Hiscock and Millero, 2006). Also due to enhanced sulfate reduction and frequently complementing methane oxidation, TA in the order of 10–2 mol kg1 can accumulate in pore waters of marine sedimentary successions (Schulz and Zabel, 2000). Even higher total alkalinity is typical for highly alkaline salt lakes, so-called soda lakes, which receive an equivalent load of HCO3 and CO32 in excess to Mg2+ and Ca2+ from their riverine inflows. Here, TA can rise to

22

ALKALINITY

more than 10–1 mol kg1 and pH increase above 10 in the course of evaporation and subsequent Ca-carbonate precipitation while alkali cations, in particular sodium and potassium, increasingly maintain the charge balance of bicarbonate and carbonate ions (e.g., Lerman and Stumm, 1989; Kempe and Kazmierczak, 1994; Reimer et al., 2009).

Alkalinity and carbonate precipitation Alkalinity, together with any of the three other measurable properties of the carbonate system – pH, dissolved inorganic carbon (DIC or CT = [CO2] + [HCO3] + [CO32]), or PCO2 (partial pressure of CO2) – can be used to compute the remaining quantities including concentrations of

Shark Bay, August 1998

2.9

1.00

SICalcite/CI

0.95 0.90 0.85

Seawater reference

ar n to ds

la

0.80

e

on

0.75

y ke

2.4

G

2.5

Lh

2.6

ol ) Po st. lin h m p t a r in Ha eg Po el la (T rb y Ca Ba on id

2.7

Calcite saturation index

2.8

M

Total alkalinity [mmol kg–1]

TA/CI

ia

M

2.3 500

700

600

800

900

1000

0.70 1100

Chloride [mmol kg–1]

b SICalcite 1.2

TA [mmol kg–1]

8.6

Ca [mmol kg–1]

16

pH

PCO2 [matm]

a

3.8

6.0

c

Westerhöfer Bach September 2005 SICalcite

12

8.2

0.9

5.6

3.6

pH 8

7.8

0.6

3.4

5.2

4

7.4

0.3

3.2

4.8

0

7.0

0.0

3.0

4.4 190 170 150

d

m a.s.l.

TA PCO2

0

50

Ca

100 150 200 250 Distance from spring [m]

300

350

e Lake Satonda October 1993

TA [mmol kg–1] 0

20

40

35

40

–0.4 0.0 0.4 0.8 1.2

SO4 / H2S [mmol kg–1]

Salinity [‰] 30

SICalcite

60

45

0

6 12 18 24

0 wind mixed aerobic surface layer

10

10-fold supersaturation

20 Depth [m]

pycnocline

30

Sal

TA

H2S anoxic middle layer

40 50 pycnocline

60

f Alkalinity, Figure 1 (Continued)

g

70

bottom layer

SO4

SICalcite

ALKALINITY

single species such as HCO3 and CO32. From CO32 and Ca2+ concentrations (or more precise activities) saturation with respect to carbonate minerals is determined, where saturation O is given by the product of the actual ion activities (IAP = {Ca2+}  {CO32}) divided by the solubility product of the carbonate (Kmineral) at ambient temperature and pressure (Omineral = IAP/Kmineral). A commonly used alternative notation is the saturation index (SI) which is defined as log O (SImineral = log (IAP/Kmineral). The oceanic CaCO3 saturation state is primarily determined by variation of the CO32 ion concentration, because the Ca2+ concentration closely depends on salinity and does not vary substantially. In the upper ocean, CO2 removal by primary production increases CO32 concentrations resulting in an about fivefold supersaturation (SI = 0.7) with respect to calcite. On the other hand, biologically controlled calcification of calcareous plankton or reefal organisms is responsible for the lower alkalinity of about 2.22.3 mmol kg1 in surface seawater. In turn, supersaturation decreases with depth when CO2 is released by remineralization of sinking organic matter when temperature and pressure changes. In the deeper ocean, total alkalinity then increases to about 2.42.5 mmol kg1 reflecting the dissolution of sinking calcium carbonate particles in response to the lower saturation state (e.g., Broecker and Peng, 1982; Morse and Mackenzie, 1990). As one of the major factors in the oceanic carbonate system, alkalinity distribution and behavior in the oceans has thoroughly been investigated during the last decades. Dissolution-preservation patterns of biogenic carbonates in surface to deep oceans and the changes brought about by a rising atmospheric CO2 level (Feely et al., 2004) have been assessed within global ocean research programs, as well as general feedbacks between the oceanic carbonate and the global climate system. Contrary to this rising knowledge on the modern marine environment, estimates

23

of ocean alkalinity throughout geological history are rare and, in the lack of direct analytical proxies, mainly rely on model assumptions. Ocean alkalinity comparably high as modern values is suggested for the Cenozoic and late Mesozoic (Tyrrell and Zeebe, 2004; Ridgwell, 2005). During periods of the Mesozoic and Paleozoic, alkalinity about two to three times higher than recent can be assumed according to the evolution of the Phanerozoic atmospheric CO2 (Ridgwell, 2005), fluctuations in the rate of continental weathering (Locklair and Lerman, 2005), or the more widespread occurrence of anoxic basins (Kempe and Kazmierczak, 1994). Elevated alkalinity concentrations of some tenths of mmol kg1 are suggested for the Earth’s primordial ocean in favor of a slightly acidic to neutral Na-Cl-dominated chemistry under an early CO2-rich atmosphere (Grotzinger and Kasting, 1993; Morse and Mackenzie, 1998). Alternatively, even higher alkalinity has been postulated for a Precambrian alkaline “Soda Ocean” in analogy to modern Na–carbonate-dominated soda lakes (Kempe and Degens, 1985). In principle, precipitation of CaCO3 is kinetically unfavorable even at the fivefold supersaturation of the modern ocean (e.g., Morse and He, 1993). Mechanisms to overcome this kinetic inhibition evolved at the Precambrian–Cambrian boundary with the advent of controlled biomineralization that became a major regulator of carbonate precipitation since the Mesozoic proliferation of the calcareous plankton. Prior to the Cambrian, carbonate deposition was characterized by sea-floor encrustations, cement beds, and the widespread occurrence of microbialites (Grotzinger, 1990; Riding, 2000), which are attributed to calcification of biofilms and microbial mats. In the lack of direct metabolic control on carbonate production, these Precambrian deposits are thought to have been formed under the pressure of a higher supersaturated environment, possibly still accounting for the dominance

Alkalinity, Figure 1 Recent environments that sustain microbialite growth illustrated in terms of alkalinity behavior and carbonate mineral saturation. (a–c) Shark Bay in Western Australia is one of the few recent marine examples that sustains calcifying biofilms involved in the formation of agglutinated stromatolites. Here, Indian Ocean water entering the embayment and running over the Faure sill into the Hamlin Pool to the SE is concentrated upon evaporation (a, courtesy to NASA). According to the twofold increase in salinity, alkalinity, and carbonate supersaturation rises continuously (b). The famous Hamlin Pool stromatolites (c) occur in the southernmost bay where alkalinity has increased to more than 2.8 mmol kg1 and waters are nearly ten times supersaturated with respect to calcite. (d and e) Today, microbialite growth is also sustained in environments such as hardwater creeks such as the Westerho¨fer Bach (Shiraishi et al., 2008), which are commonly characterized by calcium-bicarbonate type waters. From take-up of soil CO2 and subsequent subsurface dissolution of limestone rocks, these creeks have gained already high total alkalinities ranging from 4 to 6 mmol kg1. Otherwise, at their spring sites initial pH (around 7) and carbonate saturation (SICc = 0) are low due to partial CO2 pressures much higher than the atmospheric level (PCO2 = 102 atm). Continuous physicochemical CO2-degassing along the flowpath increases pH and CO32 and therefore enforces supersaturation (d). Precipitation of CaCO3 and formation of calcareous microbialites, so-called tufa stromatolites (e), commences when supersaturation reaches a certain maximum threshold level of 10- to 15-fold (SI = 1.0–1.2) (Arp et al., 2001), and results in the subsequent decline of alkalinity and Ca2+ concentrations in the creeks. (f and g) The crater lake of Satonda Island (f), Indonesia, filled with water of nearly marine composition, represents the case of a subrecent microbialite formation obviously induced by an enormous increase of alkalinity in its anoxic deep waters (Kempe and Kazmierczak, 1990). The multiple salinity stratification of the lake is a residue of former evaporatic phases. Especially in the bottom layer, sulfate has nearly been consumed due to bacterial reduction resulting in an exceptionally high total alkalinity of more than 50 mmol kg1, sulfide concentrations of up to 12 mmol kg1, and extremely high PCO2 (more than 101 atm) (g). Alkalinity and carbonate supersaturation in the photic surface layer is determined by the seasonal mixing transfer of alkalinity from the monimolimnion top layer to the mixolimnion, degassing, rain water dilution, and evaporation (Arp et al., 2003). In the surface layer alkalinity reaches twice the value of open marine waters (4.1 mmol kg1) and the tenfold supersaturation may constitute the prerequisite for the present-day, though minor, calcification of biofilms.

24

AMBER

of microbial carbonates in the early to middle Paleozoic (Riding and Liang, 2005). Today, the occurrence of carbonate deposits comparable to ancient microbialites is confined to particular environments such as restricted evaporational marine settings, freshwater habitats like tufa stromatolite forming hardwater creeks, or alkaline salt lakes (Figure 1). Although at first glance very different, these environments in common exhibit a significantly higher alkalinity than recent ocean values and a 10–15 fold supersaturation (i.e., SI = 1–1.2) with respect to calcite. Maintaining elevated carbonate supersaturation as found in these recent environments and as suggested for certain periods in Earth’s history calls for variation in alkalinity in paleo-ocean model calculations (Locklair and Lerman, 2005). Therefore, changes in alkalinity may be considered as an underappreciated hydrogeochemical key mechanism with regard to carbonate supersaturation and carbonate deposition in its various forms through geologic times.

Bibliography Arp, G., Reimer, A., and Reitner, J., 2001. Photosynthesis-induced biofilm calcification and calcium concentrations in Phanerozoic oceans. Science, 292, 1701–1704. Arp, G., Reimer, A., and Reitner, J., 2003. Microbialite formation in seawater of increased alkalinity, Satonda Crater Lake, Indonesia. Journal of Sedimentary Research, 73, 105–127. Broecker, W. S., and Peng, T.-H., 1982. Tracers in the Sea. New York: Lamont-Doherty Geological Observatory Publishers, 690 pp. Dickson, A. G., 1981. An exact definition of total alkalinity and a procedure for the estimation of alkalinity and total inorganic carbon from titration data. Deep-Sea Research, 28A, 609–623. Dickson, A. G., 1992. The development of the alkalinity concept in marine chemistry. Marine Chemistry, 40, 49–63. DOE, 1994. In Dickson, A. G., and Goyet, C. (eds.), Handbook of Methods for the Analysis of the Various Parameters of the Carbon Dioxide System in Sea Water. Version 2. ORNL/CDIAC-74. Feely, R. A., Sabine, C. L., Lee, K., Berelson, W., Kleypas, J., Fabry, V. J., and Millero, F. J., 2004. Impact of anthropogenic CO2 on the CaCO3 system in the oceans. Science, 305, 362–366. Grotzinger, J. P., 1990. Geochemical model for Proterozoic stromatolite decline. American Journal of Science, 290-A, 80–103. Grotzinger, J. P., and Kasting, J. F., 1993. New constraints on Precambrian ocean composition. Journal of Geology, 101, 235–243. Hiscock, W. T., and Millero, F. J., 2006. Alkalinity of the anoxic waters in the Western Black Sea. Deep Sea Research Part II, 53, 1787–1801. Kempe, S., 1990. Alkalinity: the link between anaerobic basins and shallow water carbonates? Naturwissenschaften, 77, 426–427. Kempe, S., and Degens, E. T., 1985. An early soda ocean? Chemical Geology, 53, 95–108. Kempe, S., and Kazmierczak, J., 1990. Chemistry and stromatolites of the sea-linked Satonda Crater Lake, Indonesia: a recent model for the Precambrian sea? Chemical Geology, 81, 299–310. Kempe, S., and Kazmierczak, J., 1994. The role of alkalinity in the evolution of ocean chemistry, organization of living systems, and biocalcification processes. In Doumenge, F. (ed.), Past and Present Biomineralization Processes. Considerations about the Carbonate Cycle. Monaco: Bulletin de l’Institut Océanographique, no. spec. 13, pp. 61–117. Lerman, A., and Stumm, W., 1989. CO2 storage and alkalinity trends in lakes. Water Research, 23, 139–146.

Locklair, R. E., and Lerman, A., 2005. A model of Phanerozoic cycles of carbon and calcium in the global ocean: evaluation and constraints on the ocean chemistry and input fluxes. Chemical Geology, 217, 113–126. Morse, J. W., and He, S., 1993. Influences of T, S, and PCO2 on the pseudo-homogeneous precipitation of CaCO3 from seawater: implications for whiting formation. Marine Chemistry, 41, 291–297. Morse, J. W., and Mackenzie, F. T., 1990. Geochemistry of sedimentary carbonates. Developments in Sedimentology. Amsterdam: Elsevier, vol. 48, 707 pp. Morse, J. W., and Mackenzie, F. T., 1998. Hadean ocean carbonate geochemistry. Aquatic Geochemistry, 4, 301–319. Rakestrow, N. W., 1949. The conception of alkalinity or excess base of seawater. Journal of Marine Research, 8, 14–20. Reimer, A., Landmann, G., and Kempe, S., 2009. Lake Van, eastern Anatolia, hydrochemistry and history. Aquatic Geochemistry, 15, 195–222. Ridgwell, A., 2005. A mid Mesozoic revolution in the regulation of ocean chemistry. Marine Geology, 217, 339–357. Riding, R., 2000. Microbial carbonates: the geological record of calcified bacterial-algal mats and biofilms. Sedimentology, 47, 179–214. Riding, R., and Liang, L., 2005. Geobiology of microbial carbonates: metazoan and seawater saturation state influences on secular trends during the Phanerozoic. Palaeogeography, Palaeoclimatology, Palaeoecology, 219, 101–115. Schulz, H. D., and Zabel, M., 2000. Marine Geochemistry. Berlin, Heidelberg: Springer, 455 pp. Shiraishi, F., Reimer, A., Bissett, A., de Beer, D., and Arp, G., 2008. Microbial effects on biofilm calcification, ambient water chemistry and stable isotope records (Westerhöfer Bach, Germany). Palaeogeography, Palaeoclimatology, Palaeoecology, 262, 91–106. Stumm, W., and Morgan, J. J., 1996. Aquatic Chemistry: Chemical Equilibria and Rates in Natural Waters, 3rd edn. New York: Wiley, 1022 pp. Tyrrell, T., and Zeebe, R. E., 2004. History of carbonate ion concentration over the last 100 million years. Geochimica et Cosmochimica Acta, 68, 3521–3530. Wolf-Gladrow, D. A., Zeebe, R. E., Klaas, C., Körtzinger, A., and Dickson, A. G., 2007. Total alkalinity: the explicit conservative expression and its application to biogeochemical processes. Marine Chemistry, 106, 287–300. Zeebe, R. E., and Wolf-Gladrow, D. A., 2001. CO2 in Seawater: Equilibrium, Kinetics, Isotopes. Elsevier Oceanography Series, Amsterdam: Elsevier, Vol 65, 346 pp.

Cross-references Calcite Precipitation, Microbially Induced Carbonate Environments Carbonates Saline Lakes Salinity History of the Earth's Ocean Soda Ocean Hypothesis

AMBER Eugenio Ragazzi1, Alexander R. Schmidt2 1 University of Padova, Padova, Italy 2 University of Göttingen, Göttingen, Germany

Synonyms Fossil resin; Resinite

AMBER

Definition Fossilized plant resin from various botanical sources. Introduction Amber is a fossilized plant resin from various botanical sources and of different geological ages, ranging from the Carboniferous to the Pliocene, although it is particularly frequent in the Cretaceous and in the Eocene to Miocene. The term “amber” is used most commonly as a synonym for fossil resin of every provenance, geological, and palaeobotanical source. A more reductive meaning of amber has been considered, referring only to succinite, the mineralogical species occurring in the Baltic coast deposits and neighboring localities containing significant amounts of succinic acid (up to 8%). However, most commonly, amber and fossil resin are the terms used interchangeably for indicating macroscopic material (Anderson and Crelling, 1995; Langenheim, 2003). Another term used by coal geologists for fossil resins is resinite, which is a microscopic material often found in coal maceral. Amber, fossil resin, and resinite, according to Anderson and Crelling (1995) should therefore be considered “solid, discrete organic material found in coals and other sediments as macroscopic or microscopic particles, which are derived from the resins of higher plants.” The term amber derives from the Arabic word anbar, which was first used to indicate ambergris (a waxy substance secreted by the sperm whale, used in perfumery as a fixative) and then also fossil resin, both natural products with the common feature of being carried ashore by sea waves. The Romans called amber succinum (from succus, literally “juice,” “sap”). The ancient Greek name for amber is elektron, that means “shining thing,” a term which was related to the Sun God Elektor (the awakener), and the Moon, called Elektris (Stoppani, 1886). The modern term electron was coined in 1891 by the Irish physicist George Johnstone Stoney, using the Greek word for amber because static electricity could be generated on the surface of rubbed amber. The German word Bernstein (or Börnstein), the Dutch Barnsteen, and the Swedish bärnsten mean “burning stone” because amber may burn when exposed to flame. Geological names are often attributed to fossil resins on the basis of the locality of discovery or of the discoverer, and these names are not based on specific knowledge of the composition; however, they are often used and also quoted in the scientific literature as common names. For example, Simetite is for amber collected in Sicily (Italy) near the Simeto river, Burmite, for amber found in Burma (Myanmar), and Rumanite for that found in Romania; Beckerite and Stantienite were named for Becker and Stantien, two developers of amber dredging and mining operations in the Samland region in the 1800s. Over a hundred of different fossil resins have been described, so far. Fossil resins are considered the archetype of a “chemical fossil” (chemical evidence of life processes preserved in the fossil record), and besides their property of

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preserving embedded organisms, they can also maintain particular details of their original composition better than any other sedimentary material. Amber has been known by humans since prehistorical times and most bizarre hypotheses about its origin and composition have been formulated. In the antiquity amber has been considered even as produced by the rays of the Sun (Pliny the Elder, Book 37: Nicias solis radiorum sucum intellegi voluit hoc) and that was not such a wrong idea, since amber, as a resin of ancient trees, is a metabolite of plants, originated from products of photosynthesis, that is, from the energy of the Sun.

Amber composition and formation Several different parameters can play relevant roles in amber composition and consequently, they all should be considered in the study and comparison of different amber samples. These are for example: 1. Original environment: temperature, insolation, and climate influence the resin composition before it is embedded in the sediment (biostratonomic processes); 2. Age: complex chemical arrangements occur with time; in general, low molecular weight components disappear with increasing age; 3. Plant source: resin produced by different plants exhibit different chemical composition; 4. Diagenetic history: following the embedding in the sediment, resin maturation is affected by temperature, pressure and permeating fluids which can modify the chemical composition and the physicochemical properties of fossil resin; 5. Alteration processes after amber mining (of relevance especially in archaeological context): direct sunlight and exposition to air and heat may change the color and physical properties (and, consequently, the chemical composition) of the sample. Resins consist of secondary components of plants, that is, chemicals occurring only in some groups of plants, without an apparent role in primary physiological or metabolic pathways. Most natural resins are derived from terpenes, a large and varied class of hydrocarbons, which are chemicals composed by structures based on the isoprene unit (C5H8) (Figure 1). The basic molecular formula of terpenes can be expressed as multiples of isoprene, (C5H8)n, where n is the number of linked isoprene units. The isoprene units can link together “head to tail” to form linear chains, or alternatively they can arrange to form rings. The overall terpene structure undergoes oxidation or rearrangement of the carbon skeleton, producing compounds, which are generally indicated as terpenoids or isoprenoids. Mono-, sesqui-, di-, and tri-terpenoids are polymers formed by two, three, four, and six isoprene units, respectively. The biosynthesis of resins starts from simple molecules, namely mevalonic acid, which can be considered the precursor of isoprene units. Photosynthetically

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AMBER

produced carbohydrates are broken down to give pyruvate, which represents the basis for the synthesis of mevalonic acid. Mevalonic acid is converted into the activated form of isopentenyl pyrophosphate, which gives origin to geranyl pyrophosphate (C10), the basis for the monoterpene synthesis, or can further undergo condensation into farnesyl pyrophosphate (C15) for the production of sesquiterpenes, or geranyl-geranyl pyrophosphate (C20), which yields diterpenes and tetraterpenes, or squalene (C30), which is the basis of triterpenes (Figure 2). Fluid resins, after their exudation from plant, are characterized by a large volatile fraction, up to 50%, mainly constituted by monoterpenes and sesquiterpenes, while the nonvolatile fraction is composed of di- and tri-terpene acids (carboxylic acids with double bonds and functional groups), with alcohols, aldehydes, and esters. The volatile components of the resins are mainly involved in the defensive role of the resin against pathogens and insects. Evaporation of volatile components, following polymerization of the remaining nonvolatile constituents, mainly presenting dienic functions, causes the hardening of resin, which, if allowed to be embedded in appropriate sediment, poses the basis for transformation into a fossil resin.

Amber, Figure 1 The basic isoprene unit (C5H8).

Diterpenoid resins, mainly composed of diterpene acids (C20), are produced by gymnosperms (conifers) and angiosperms (mostly in the Leguminosae family), and they are particularly prone to polymerization and thus being preserved during fossilization into amber. Gymnosperm diterpenes are mainly represented by diterpene acids of the abietane, pimarane, and labdane types (Figure 3). Abietane- and pimarane-type diterpene acids, such as abietic and pimaric acids, abundant in the Pinaceae family, tend to remain unpolymerized; on the contrary, labdane-type acids (e.g., agathic and communic acids, common in the Cupressaceae family), easily polymerize thanks to conjugated diene compounds (Langenheim, 2003). Resins of the Araucariaceae family may contain all three diterpene groups. Also the nonvolatile fraction of angiosperm resins, for example, from the genus Hymenaea of the Leguminosae (Fabaceae) family, which yields copal, is constituted by diterpenes of the labdadiene group. Resins from other angiosperms, such as Dipterocarpaceae, contain mainly triterpenes. Therefore, resins containing particular diterpenes, such as abietic acid, cannot form amber sensu stricto because the constituent does not contain functional groups that could lead to a cross-linked polymer (Beck, 1999). Also triterpenoid resins (C30), which are mainly produced by broad-leaved trees, generally do not polymerize. The process of polymerization and resin hardening starts just after resin exudation, and involves free radical mechanisms, which are initiated by exposure to sunlight and air (Langenheim, 2003). Initial polymerization can

Amber, Figure 2 Biosynthesis of terpenoid resins from mevalonate (adapted from Langenheim, 1969).

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Amber, Figure 3 Chemical structure of major conifer resin components. Left column presents the general skeleton, and right column, an example of specific compounds (for further details, see Langenheim, 2003).

occur involving the terminal groups of a side chain in the molecule of terpenoids, mainly diterpenes with a labdanoid structure (unsaturated labdatriene acids and alcohols, and biformene), leading to the formation of a general polymeric structure (Anderson et al., 1992; Clifford and Hatcher, 1995; Clifford et al., 1997) (Figure 4). As a complex mixture, amber contains also nonpolymerizing compounds such as succinic acid in succinite, or monoterpenes, which are trapped (cross-linked) into the polymer network (Clifford and Hatcher, 1995; Clifford et al., 1997) and function as plasticizers. Formation of amber occurs through the process of “maturation,” which is defined as the progressive changes of the resin occurring after the hardening and burial into the sediment, due to diagenetic and catagenetic processes. The chemical reactions in maturation of the resin to yield fossil resin include cross-linking, isomerization, and cyclization (Clifford and Hatcher, 1995; Clifford et al., 1997). One typical reaction in polylabdanoid fossil resins is the depletion of exocyclic methylenes (Figure 4) in the C8–C17 double bond of the molecule, and the formation of intramolecular cyclization. The modification of amber structure during its maturation leads to a progressive loss of unsaturated bonds, a decrease in functionalized groups and an increased proportion of aromatized groups (Grimalt et al., 1988).

Although the degree of resin maturation is dependent on the age of the resin, the rate at which this process occurs depends on several geological conditions, including the thermal history of the sample (Anderson et al., 1992) and on the structure and resin composition (Langenheim, 2003). Although a certain direct correlation between resin maturation and the age has also been demonstrated, by means of thermal analysis (Ragazzi et al., 2003), maturity and age cannot always be directly linked, since chemical transformation of resin increases at higher temperatures, and therefore the geothermal variable should be considered. Other physicochemical properties of amber, such as hardness and solvent solubility, may suggest the degree of maturation (Poinar, 1992; Rodgers and Currie, 1999; Langenheim, 2003; Ragazzi et al., 2003). An alternative approach to define the age of a resin was proposed by Anderson (1997) on the basis of carbon-14 content. Resins that are older than the detection limit of carbon-14, that is, at about 40,000 years, have been considered fossil resins, and their age should be determined based on the stratigraphical data of the embedding sediment. Resins of age between 40,000 and 5,000 years are considered subfossil, while resins between 5,000 and 250 years are ancient resins, and those younger than 250 years are considered as modern or recent. This

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Amber, Figure 4 Schematic representation of the polymeric structure in polylabdanoid fossil resins and of the maturation process (adapted from Clifford and Hatcher, 1995; Clifford et al., 1997).

terminology should be preferred to that referring to “young amber” or “copal,” which is often misleading with respect to the age of the sample.

Physicochemical characteristics of amber Most amber pieces are minute ranging from millimeter to centimeter size. Resin pieces may be found preserved in their original shape as solidified in the amber forest (e.g., drops, elongate stalactites, lumps with a flattened side which was attached to the bark or to other surfaces). Fossil resin is sometimes still attached to fossil wood, or even in situ in wood or twigs which can help to identify the amber-bearing trees of a particular deposit. Amber pieces are more or less rounded after being transported and redeposited. Amber has been extensively investigated under the physicochemical aspect. The relative hardness according to Mohs’ scale is 2–2.5 (corresponding to 199–290 MPa) for most types of amber, indicating that it is slightly harder than gypsum and that it is possible to scratch the surface with calcite or copper. The specific

gravity, that is, the ratio of the density of a substance to the density of water, ranges from about 0.96–1.12. For this reason most amber is nearly floating in sea water (specific gravity of the Baltic sea water is very low, about 1.005) and it sinks to the bottom in quiet water, but it can easily be carried along by disturbed water and washed ashore. Because of its low density, amber floats on a salt solution (about 15 g of NaCl/100 ml), while several types of plastic, often used as amber imitation, sink due to their higher density (e.g., the phenolic plastic bakelite, which has a specific gravity of about 1.4). Amber has resinous luster, and might appear translucent or opaque. Its refractive index ranges between 1.539 and 1.545. The color of translucent amber varies from a honey-like yellow and yelloworange to light and dark red; the latter color is often caused by weathering. Rare specimens of blue (e.g., some Dominican amber) and green (e.g., some Mexican amber) samples have been found. Resins may be opaque because of microscopic bubbles or tiny detritus particles. Sometimes, densely arranged filaments of bacteria and fungi

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that grew into the resin in its liquid stage cause opacity of the outer parts of resin pieces (Figure 5). If exposed to ultraviolet light, amber often produces fluorescence of blue, yellow, or green color. When amber is heated, it becomes soft at 150–180 C and decomposes around 280–300 C, emitting fumes with aromatic odor. After touching amber with a hot needle (hot point test), it produces smoke and burnt-resinous odor; this method is empirically used to distinguish between amber and plastic or copal. Exposed to a flame, amber burns easily. Amber is insoluble in water, and partly soluble in organic solvents,

Amber, Figure 5 Filamentous sheathed bacteria of ca 10 mm diameter preserved in French Cretaceous amber after growing into the liquid resin (photograph by Alexander R. Schmidt).

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such as diethyl ether, acetone, and dichloromethane, as well as in mixtures of turpentine oil and acetone. After applying a drop of solvent to the surface, recent resins (copal) or plastic are readily dissolved and the surface becomes sticky, while amber (polymerized fossil resin) is unaffected. Although resin is macroscopically amorphous, some of its constituents may form microscopic drop- or bubble-like and elongated microstructures. Some of these “pseudoinclusions” may resemble enclosed microorganisms but can be distinguished by their variable size and shape and by the absence of diagnostic surface features (Figure 6).

Chemical analyses and classification of amber Since early chemical investigations in the nineteenth century, amber has been identified as being composed of carbon, hydrogen, and oxygen. It is not possible to indicate a univocal composition formula of amber since it is an amorphous, noncrystalline material. The composition of different types of amber, as a result of elemental analysis, has been indicated approximately in the following range: carbon 75–87%, hydrogen 8.5–11%, oxygen 5–15%, with traces of sulfur (up to 1.7%) and other elements (Kosmowska-Ceranowicz, 1984; Ragazzi et al., 2003). Since amber is mostly insoluble in organic solvents, a complete characterization of its chemical composition has been obtained using the total solid sample, by means of specific analytical techniques that have become available, such as infrared spectroscopy (IRS), pyrolysis-gas chromatography/mass spectrometry (Py-GC/MS), and nuclear magnetic resonance (NMR) spectroscopy.

Amber, Figure 6 Pseudoinclusions in Cenomanian amber from Fourtou in the French Pyreneans (photograph by Alexander R. Schmidt).

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AMBER

Infrared spectroscopy of powdered samples, since the first application in the study of fossil resins (Beck et al., 1964), has become one of the main techniques for obtaining structural information and identification of amber from different origins (Kosmowska-Ceranowicz, 1999). Infrared spectra allow to define the fingerprinting of any fossil resin, and in particular it has been successful as diagnostic for Baltic amber, observing the presence of the characteristic Baltic shoulder in the 1,150– 1,260 cm1 region of the spectrum, attributed to the absorption of ester groups, in particular esterified succinic acid. It is noteworthy that the free succinic acid levels in Baltic amber range between 50 and 400 ppm (Tonidandel et al., 2009), despite the high total succinic acid content of 1–8%. Anderson et al. (1992; also Anderson and Crelling, 1995), on the basis of various physicochemical methods, mainly Py-GC/MS, proposed a classification of fossil resins into five main classes: Class I

Class II

Class III Class IV Class V

The macromolecular structures are derived from polymers of primarily labdanoid diterpenes, which include labdatriene carboxylic acids, alcohols, and hydrocarbons Ia Derived from/based on polymers and copolymers of labdanoid diterpenes with regular [1S, 4aR, 5S, 8aR] configuration, including communic acid, communol, and succinic acid; example: succinite (Baltic amber) Ib Derived from/based on polymers and copolymers of labdanoid diterpenes with regular [1S, 4aR, 5S, 8aR] configuration, often including communic acid, communol, biformene, but not succinic acid; example: New Zealand’s fossil kauri resin Ic Derived from/based on polymers and copolymers of labdanoid diterpenes with enantio [1S, 4aS, 5R, 8aS] configuration, including ozic acid, ozol, and enantio biformenes; examples: Mexican and Dominican amber, African resinites Derived from/based on polymers of bicyclic sesquiterpenoid hydrocarbons, especially cadinene, and related isomers; examples: Utah and Indonesian amber Natural (fossil) poylstyrene; example: some New Jersey amber, Siegburgite from Germany Nonpolymeric, incorporating sesquiterpenoids based on the cedrane carbon skeleton; example: Moravian amber from the Czech Republic Nonpolymeric diterpenoid carboxylic acid, especially based on abietane, pimarane, and isopimarane carbon skeletons; example: Highgate copalite and other retinites in European brown coals

The high-resolution NMR spectra of the carbon nuclei in powdered samples provided detailed information on the types of carbon functionality in the fossil resin, over the entire range of resonances, indicating the presence of

saturated carbons (resonances 0–100 ppm), unsaturated carbons such as alkenic and aromatic (resonances 100–160 ppm), and carbonyl groups (resonances above 160 ppm) (Lambert and Frye, 1982). Using NMR, comparative analysis of samples from different origin permitted to distinguish four major groupings (Group A through D) of fossil resins (Lambert and Poinar, 2002), in good agreement with Anderson’s et al. classification based on Py-GC/MS.

Age, global occurrence, and botanical origin of amber Amber might be found on every continent but most amber deposits have been discovered in the northern hemisphere, so far. A few ambers are known from South America, Africa, Australia, and New Zealand. Many new Mesozoic and Cenozoic amber localities have been described in the last decade (see Poinar, 1992; Grimaldi, 1996; Krumbiegel and Krumbiegel, 2005, for age and global occurrence of ambers). A few resins were reported from the Carboniferous of Europe and the USA (Bray and Anderson, 2009). The first conifers which produced significant amounts of resin appeared in the Carnian stage of the late Triassic (about 220 million years ago). Remarkable amounts of small Triassic amber pieces have been found in the petrified forests of Arizona (Litwin and Ash, 1991), in the Molteno Formation of Lesotho (Ansorge, 2007) and the HeiligkreuzSanta Croce Formation of the Italian Dolomites (Roghi et al., 2006, Figure 7). The latter is the oldest fossiliferous amber containing various microorganisms (Schmidt et al., 2006, see Figure 8). The Jurassic is particularly poor in amber and just few samples have been reported from the late Jurassic age of Portugal, Denmark (Bornholm), Germany, and Thailand. The oldest Cretaceous ambers come from the Wealden Formation of southern England (Nicholas et al., 1993) and from the Kirkwood Formation of South Africa (Gomez et al., 2002), both being Valanginian in age. The most important fossiliferous Cretaceous ambers are (1) the Middle East ambers from Lebanon, Israel, and Jordan, ranging from the Hauterivian to the Aptian, (2) the Aptian–Albian Àlava amber from Peñacerrada in the Basque county of northern Spain, (3) the Siberian amber from the Albian to Santonian of the Taimyr Peninsula, (4) the Albian–Cenomanian ambers from Charente and Charente-Maritime in southwestern France (see Perrichot et al., 2007, for these newly discovered ambers), (5) the Burmese amber from Myanmar which is Albian in age (Grimaldi et al., 2002; Cruickshank and Ko, 2003), (6) the Atlantic coastal plain ambers of the USA which comprise several localities of various Upper Cretaceous ages from Maryland and New Jersey to South Carolina, and (7) the Upper Cretaceous Canadian amber (Cedar Lake in Manitoba and other localities). Further remarkable Cretaceous resins come from Alaska, from several Albian to Coniacian localities in Japan, and from

AMBER

Amber, Figure 7 Triassic amber drop from the Italian Dolomites (photograph by Guido Roghi, Nature, 444, 835, 2006). The piece is about 3 mm in length.

Amber, Figure 8 Shell of a testate amoeba enclosed in Triassic amber from Italy. This shell is 35 mm in size and morphologically identical to the modern species Centropyxis hirsuta (photograph by Alexander R. Schmidt, Nature, 444, 835, 2006).

the Aptian age of Golling in Austria, but these ambers yield few fossils. Cenozoic ambers are largely Eocene to Miocene in age, and some deposits already produced tons of resin. The most well-known fossil resin is the Baltic amber which

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can be found in the Baltic sea area (especially along the coast of Russia and Poland) in Eocene sediments, washed ashore and also as glacial erratic. The sediments bearing the Baltic amber are considered to be 38–47, some up to 55 million years old (Ritzkowski, 1999). Other fossiliferous Eocene ambers come from the Paris Basin in France (50–55 million years old; Nel et al., 1999), from Vastan (Gujarat) in western India (52 million years old; Alimohammadian et al., 2003) and from Fu Shun in the Chinese Liaoning province. The Bitterfeld amber from a coal mine near the city of Bitterfeld in central Germany is uppermost Oligocene in age (24 million years old, see Wimmer et al., 2006) and also produced a plethora of inclusions, mainly arthropods (Knuth et al., 2002). The most fossiliferous Cenozoic New World ambers come from the Dominican Republic and from Chiapas in Mexico (see Poinar, 1992). Recent studies suggest a Miocene age (15–20 million years old) for these Caribbean ambers (Iturralde-Vinent and MacPhee, 1996; Solórzano Kraemer, 2007). Further fossiliferous Miocene ambers were found in western Amazonia in Peru, and in Sicily and Borneo. Representatives of various gymnosperm and angiosperm families have produced resin in the Earth’s history (see Langenheim, 2003; Krumbiegel and Krumbiegel, 2005, for overview). The Italian Triassic amber was possibly produced by members of the Cheirolepidiaceae, an extinct gymnosperm family. Among others, various representatives of the Araucariaceae (Figures 9 and 10) and Cupressaceae are considered to be resin-bearing trees in the Cretaceous. Angiosperm resins are recorded since the Eocene, for example, Glessite stems from Burseraceae and Siegburgite were produced by Hamamelidaceae. The tropical genus Hymenaea (Leguminosae) produced the Dominican and Mexican ambers. The main resin-bearing trees of the Baltic amber forest probably belonged to conifers of the family Sciadopityaceae, as recently suggested by Wolfe et al. (2009).

Embedding and preservation of organisms in resin Resins are produced by plants as defensive reaction against mechanical damage (injury to the bark, fire, herbivores) and potential invasion by pathogens, such as bacteria and fungi. Also, climatic reasons and ecological disturbance have been discussed to be the reasons for extensive resin production (Henwood, 1993). The exceptional preservation of organisms included in amber is due to the dehydration environment made possible by the resin and by natural embalming properties due to antiseptic activity of the resin constituents. Components that do not participate in polymerization of the resin, such as some diterpene resin acids (e.g., copalic acid), can remain available for the interactions with biota embedded in resin (Henwood, 1993) and contribute to the preservation even of soft-bodied inclusions. Even cell organelles such as chloroplasts of algae are sometimes wellpreserved in amber (Figure 11).

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Amber, Figure 9 Resinous forest of Araucaria columnaris growing at the coast of Mare´, New Caledonia (photograph by Alexander R. Schmidt).

Amber, Figure 11 Preservation of chloroplasts in the conjugatophyte Palaeozygnema spiralis from the Cenomanian resin of Schliersee in the Bavarian Alps. The cells are 15 mm in diameter (photograph by Alexander R. Schmidt).

Amber, Figure 10 Extensive resin flows of 30 cm length induced by fire at the bark of Araucaria columnaris in New Caledonia (photograph by Alexander R. Schmidt).

Extraction of ancient DNA from amber has been of particular interest in working with fossils embedded in amber, and in the early 1990s first attempts to establish methods for extraction, amplification, and sequencing of fossil

DNA from amber were carried out (e.g., Cano et al., 1994; Cano and Borucki, 1995). However, critical reinvestigations suggest that the DNA obtained in these studies was modern contamination and because even the amplification of DNA from copal failed, Austin et al. (1997) concluded that resin is not predestined for the preservation of complex molecules, although morphological structures are recorded very well.

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Attached to fresh sticky resin (Figure 10), organisms or parts of them are entirely covered by further resin flows. Amber inclusions are therefore usually found at the surfaces of successive resin flows in amber. Invertebrates of appropriate size, especially arthropods (e.g., diptera, hymenoptera, see Figure 12), and small plant remnants fallen onto the resin are therefore predestined for getting stuck and embedded. Arthropods, plant remnants (flowers, leaves, stellate hairs, wood, pollen, spores) and microorganisms are typical amber inclusions and sometimes, also small vertebrates (frogs, lizards) and parts of larger animals (reptile skin, feathers, hairs) are preserved. Some groups of organisms are only recorded as amber fossils, since few other fossilization processes occur in terrestrial environments which may preserve morphological structures even of soft-bodied organisms very well. Many life forms and representatives of all trophic levels of a biocoenosis may be found in a single piece of resin: bacteria, algae as producers, protozoans, micrometazoans, molluscs, arthropods as consumers, and fungi as decomposers (Perrichot and Girard, 2009). Although many members of a biocoenosis can potentially become embedded in resin, selective embedding has been observed (Schmidt and Dilcher, 2007). Because of their motility, moving or flying arthropods have a higher probability of encountering and becoming attached to the resin. Therefore, the amount of amber inclusions of a species is not correlated with its abundance in the ancient ecosystem. Most amber inclusions are fossils of terrestrial organisms which lived at or close to the resin-bearing trees. Much resin solidified at the forest floor, not on the bark (Henwood, 1993) and the term litter amber has been proposed by Perrichot (2004) for amber pieces which largely contain soil and litter-dwelling arthropods (Figure 13).

To find aquatic organisms in tree resin may seem to be highly unlikely but the fossil record provides amber-preserved limnetic arthropods (e.g., water beetles, water striders, crustaceans, larvae of caddisflies and mayflies) and microorganisms (e.g., algae, ciliates, testate amoebae, rotifers). Limnetic organisms may get stuck or enclosed when resin comes into contact with water or even flows into water (Schmidt and Dilcher, 2007). Few amber inclusions of marine organisms (crustaceans, diatoms) have been reported from amber forests which grew directly at the coast (Vonk and Schram, 2007; Girard et al., 2008; see Figure 14). These organisms were probably introduced by wind or spray from beach and sea onto the resin flows in the nearby woods. Although resin has fungicidal properties, some fungi and bacteria are able to grow into liquid resin and can therefore be found in almost every fossil resin. Usually, these filaments grow in random orientation as long as the resin is liquid. Growth stops when the resin solidifies and the filiform inclusions become well-preserved in the resin. Resinicolous fungi such as ascomycetes of the Mycocaliciales are adapted to their special substrates and are also able to grow on fresh resin. Therefore, they may be found in places where herbivore insects induce longterm resin flows (Rikkinen, 1999; Rikkinen and Poinar, 2000). Resin which dried at the bark is exposed to the air, and processes of weathering and degradation start rapidly. Long-time preservation of resins and its inclusions and

Amber, Figure 12 Hymenoptera (Scelionidae), fossilized in Lower Cenomanian amber of Fouras, Charente-Maritime, southwestern France. The inclusion is 1.8 mm in length (courtesy of Vincent Perrichot).

Amber, Figure 13 Marchandia magnifica, a mole cricket from Albian amber from southwestern France. It is 3.8 mm in length (photograph by Vincent Perrichot, courtesy of Elsevier/ Academic Press, Cretaceous Research, 23, 307–314, 2002).

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AMBER

Amber, Figure 14 Two frustules of a marine diatom of the genus Hemiaulus enclosed in a piece of amber from a coastal Cretaceous forest from Archingeay in south-western France. The cells are ca 40 mm in size (photograph by Alexander R. Schmidt).

formation of amber therefore generally needs redeposition into marine sediments or at least a cover by sediment layers to stop processes of oxidation.

Summary Amber is fossilized plant resin, which is largely found in the Cretaceous and in the Eocene to Miocene when various representatives of gymnosperms and angiosperms produced considerable amounts of resin. After exudation from plants, the volatile components evaporate and the remaining nonvolatile constituents polymerize which leads to the hardening of the resin. Formation of amber occurs through the process of “maturation,” which is defined as the progressive changes of the resin occurring after the hardening and embedding into the sediment. The chemical reactions in the maturation of the resin to yield fossil resin include cross-linking, isomerization, and cyclization. Several different parameters can play relevant roles in amber composition, such as original environment, age, plant source, diagenetic history, and also alteration processes after amber mining. Inclusions of many life forms may be found in amber, although selective embedding has been observed. The taphocoenosis in a piece of amber may potentially contain organisms of different habitats such as tree bark, soil and litter, freshwater ponds, and littoral.

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M., Bignot, G., Cavagnetto, C., Duffaud, S., Gaudant, J., Hua, S., Jossang, A., De Lapparent de Broin, , F., Pozzi, J. P., Paicheler, J. C., Bouchet, F., and Rage, J. C., 1999. Un gisement sparnacien exceptionnel à plantes, arthropodes et vertébrés (Éocène basal, MP7): Le Quesnoy (Oise, France). Comptes Rendus de l’Académie des Sciences, Sciences de la Terre et des Planètes, 329, 65–72. Nicholas, C. J., Henwood, A. A., and Simpson, M., 1993. A new discovery of Early Cretaceous (Wealden) amber from the Isle of Wight. Geological Magazine, 130, 847–850. Perrichot, V., 2004. Early Cretaceous amber from south-western France: insight into the Mesozoic litter fauna. Geologica Acta, 2, 9–22. Perrichot, V., and Girard, V., 2009. A unique piece of amber and the complexity of ancient forest ecosystems. Palaios, 24, 137–139. Perrichot, V., Néraudeau, D., Nel, A., and De Ploëg, G., 2007. A reassessment of the Cretaceous amber deposits from France and their palaeontological significance. African Invertebrates, 48, 213–227. Poinar, G. O., Jr., 1992. Life in Amber. Palo Alto: Stanford University Press. Ragazzi, E., Roghi, G., Giaretta, A., and Gianolla, P., 2003. Classification of amber based on thermal analysis. Thermochimica Acta, 404, 43–54. Rikkinen, J., 1999. Two new species of resinicolous Chaenothecopsis (Mycocaliciaceae) from western North America. The Bryologist, 102, 366–369. Rikkinen, J., and Poinar, G. O., Jr., 2000. A new species of resinicolous Chaenothecopsis (Mycocaliciaceae, Ascomycota) from 20 million year old Bitterfeld amber, with remarks on the biology of resinicolous fungi. Mycological Research, 104, 7–15. Ritzkowski, S., 1999. Das geologische Alter der bernsteinführenden Sedimente in Sambia (Bezirk Kaliningrad), bei Bitterfeld (Sachsen-Anhalt) und bei Helmstedt (SE-Niedersachsen). In: Investigations into Amber. Proceedings of the International Interdisciplinary Symposium: Baltic Amber and Other Fossil Resins, September 2–6, 1997, Gdansk. The Archaeological Museum in Gdansk, Museum of the Earth, Polish Academy of Sciences, Gdansk, pp. 33–40. Rodgers, K. A., and Currie, S., 1999. A thermal analytical study of some modern and fossils resins from New Zealand. Thermochimica Acta, 326, 143–149. Roghi, G., Ragazzi, E., and Gianolla, P., 2006. Triassic amber of the southern Alps (Italy). Palaios, 21, 143–154. Schmidt, A. R., and Dilcher, D. L., 2007. Aquatic organisms as amber inclusions and examples from a modern swamp forest. Proceedings of the National Academy of Sciences of the United States of America, 104, 16581–16585. Schmidt, A. R., Ragazzi, E., Coppellotti, O., and Roghi, G., 2006. A microworld in Triassic amber. Nature, 444, 835. Solórzano Kraemer, M. M., 2007. Systematic, palaeoecology, and palaeobiogeography of the insect fauna from the Mexican amber. Palaeontographica Abteilung A, 282, 1–133. Stoppani, A., 1886. L’ambra nella storia e nella geologia, con speciale riguardo agli antichi popoli d’Italia nei loro rapporti colle origini e collo svolgimento della civiltà in Europa. Milan, Italy: Dumolard. Tonidandel, L., Ragazzi, E., and Traldi, P., 2009. Mass spectrometry in the characterization of ambers. II. Free succinic acid in fossil resins of different origins. Rapid Communication in Mass Spectrometry, 23, 403–408. Vonk, R., and Schram, F. R., 2007. Three new tanaid species (Crustacea, Peracarida, Tanaidacea) from the Lower Cretaceous Álava amber in Northern Spain. Journal of Paleontology, 81, 1502–1509.

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Wimmer, R., Pester, L., and Eissmann, L., 2006. Das bernsteinführende Tertiär zwischen Leipzig und Bitterfeld. Mauritiana (Altenburg), 19, 373–421. Wolfe, A. P., Tappert, R., Muehlenbachs, K., Boudreau, M., McKellar, R. C., Basinger, J. F., and Garrett, A., 2009. A new proposal concerning the botanical origin of Baltic amber. Proceedings of the Royal Society B, 276, 3403–3412.

Cross-references Algae (Eukaryotic) Bacteria Biological Control on Diagenesis: Influence of Bacteria and Relevance to Ocean Acidification Biomarkers (Molecular Fossils) Diatoms Fungi and Lichens Geomycology Protozoa (Heterotroph, Eukaryotic)

ANAEROBIC OXIDATION OF METHANE WITH SULFATE Katrin Knittel, Antje Boetius Max Planck Institute for Marine Microbiology, Bremen, Germany

Definition Anaerobic oxidation of methane (AOM): microbially mediated oxidation of methane to CO2 by electron acceptors other than oxygen. Introduction Methane is the most abundant hydrocarbon in the atmosphere, and an important greenhouse gas (see Methane, Origin). A great deal of research has focused on the cause and climatic consequences of the variation in fluxes of methane to the atmosphere, throughout the Earth’s history. Three key functional groups of microbial organisms play a central role in regulating the fluxes of methane on the Earth, namely the methanogens, the aerobic methanotrophic bacteria, and the more recently discovered anaerobic methanotrophic archaea (ANME). It is estimated that AOM is a major sink for methane on the Earth, and of similar relevance as its photooxidation in the atmosphere (Hinrichs and Boetius, 2002; Reeburgh, 2007). Today, most methane is produced by methanogenesis, i.e., the final step in the fermentation of organic matter taking place in soils, wetlands, landfills, rice fields, freshwater and marine sediments, as well as in the guts of animals. Almost all of the methane produced in ocean sediments is consumed by AOM within the sulfate penetrated seafloor zones. Hence, the ocean does not contribute significantly to the atmospheric methane budget (40 µm in diameter (Nauhaus et al., 2007; Holler, unpublished data). After reaching a specific size they appear to burst, releasing single cells into the environment (Figure 2ab). The ANME-3 clade also belongs to the order Methanosarcinales and is closely related to cultivated species of the genus Methanococcoides with 95% 16S rRNA sequence similarity (Figure 1a). They form shelltype aggregates with a Desulfobulbus population as sulfate-reducing partner, however, only very few bacterial cells are associated (Figure 2y–z, aa).

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Anaerobic Oxidation of Methane with Sulfate, Figure 1 (Continued)

ANAEROBIC OXIDATION OF METHANE WITH SULFATE

Identification of ANME by methyl coenzyme M reductase gene phylogeny The first genomic and proteomic analyses of sediments and microbial mats naturally enriched in the ANME biomass revealed a nickel protein similar to methyl coenzyme M reductase (MCR) as well as genomic fragments coding for nearly all genes typically associated with methane production. Based on these observations, earlier hypotheses on the role of reverse methanogenesis in archaea (Zehnder and Brock, 1979, 1980; Hoehler et al., 1994) appeared in a new light. Subsequently, molecular studies were conducted targeting the enzyme MCR, which catalyzes the terminal step in biogenic methane production (Shima and Thauer, 2005). Investigations have revealed a remarkably high phylogenetic diversity within mcrA genes among ANME archaea (Hallam et al., 2003; Inagaki et al., 2004, 2006a; Dhillon et al., 2005; Kelley et al., 2005; Alain et al., 2006; Lloyd et al., 2006; Nunoura et al., 2006; Lösekann et al., 2007) which have been classified into four subgroups (Hallam et al., 2003; Lösekann et al., 2007), group a–b (ANME-1), group c–d (ANME-2c), group e (ANME-2a), and group f (ANME-3) (Figure 1b). These groups are all distinct from those formed by methanogens. ANME mcrA gene phylogeny appears to be partially phylogenetically congruent to the 16S rRNA gene (Figure 3; Hallam et al., 2003; Nunoura et al., 2006; Lösekann et al., 2007). Quantification of specific ANME groups based on their mcrA gene abundance is now an alternative method to 16S rRNA based FISH to study distributional patterns of anaerobic methanotrophs in AOM zones (Nunoura et al., 2006). Identification of ANME by their specific lipid biomarkers Another molecular method, which has been crucial in the investigation of the natural distribution of ANME populations, is the identification of specific lipid biomarkers (see Biomarkers) and their stable carbon isotope signatures. This method can integrate phylogenetic information with function (13C signatures indicating methane assimilation) and can be used for comparative analyses of community biomass. All three known ANME groups and their partner bacteria incorporate light (13C-depleted) methane-derived carbon into their membrane lipids (Orphan et al., 2002; Niemann et al., 2006b). Using either naturally 13C depleted methane (Nauhaus et al., 2007) or 13 C labeled methane as substrate (Blumenberg et al., 2004) for active populations of ANME in environmental samples have helped to identify typical membrane lipid

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profiles. Crocetane is the only biomarker lipid specific for ANME, i.e., not found in methanogen cultures. Glycerol dibiphytanyl glycerol tetraethers (GDGT) were also used as indicator of ANME distribution (Schouten et al., 2003), but the ANME GDGTs appear to show some overlap with those of benthic Crenarchaeota, which often share the same niche in the seabed (Knittel et al., 2005). Recently, the biomarker approach was extended to include intact polar lipids (IPLs) which are of higher taxonomic specificity (level of families to orders) and property to select for living biomass (Sturt et al., 2004; Biddle et al., 2006; Rossel et al., 2008). Characteristic IPL molecular fingerprints have been reported for each specific ANME type: the lipids of ANME-1 archaea were dominated by diglycosidic GDGT derivatives, while IPLs of ANME-2 and ANME-3 archaea are dominated by phosphate-based polar derivatives of archaeol and hydroxyarchaeol (Rossel et al., 2008).

Bacterial partners of ANME ANME-1 and ANME-2 archaea are usually associated with SRB of the SEEP-SRB I branch, subgroup a, of the Desulfosarcina/Desulfococcus (DSS) group (Figure 2; Schreiber et al., 2010). These SRB are physically attached to the ANME-2 archaea, whereas ANME-1 is most often found without close physical association to other cells. The diversity within the SEEP-SRB1 cluster is high, and it is still unknown if the partners of the ANME-2a, -2b, and -2c subgroups belong to the same species. The morphology of the DSS cells varies from cocci (mostly associated with ANME-2c cells, Figure 2p–x) to rods (mostly associated with ANME-2a cells, Figure 2f–j). Other SRB have not yet been identified to be associated with ANME-1 and -2. ANME-3 archaea are most often associated with SRB of the Desulfobulbus branch (Figure 2), but have also been detected together with SRB of the Desulfosarcina/ Desulfococcus group in shallow subsurface sediments of cold seeps from the Hydrate Ridge (Lösekann et al., unpublished data). Single cells of ANME-1, -2, and -3 have also been found in situ as well as in vitro enrichments, indicating that the physical association may be not obligate, but certainly the typical life mode of the anaerobic methanotrophs in most habitats. There is also some evidence that the diversity of bacteria associated with ANME may be larger than anticipated. The analysis of ANME-2c consortia captured by wholecell magneto-FISH showed a diversity of bacterial partners of ANME-2 far beyond the Deltaproteobacteria

Anaerobic Oxidation of Methane with Sulfate, Figure 1 Phylogenetic trees showing the affiliations of (a) anaerobic methanotrophic archaea (ANME) 16S rRNA gene sequences to selected reference sequences of the domain archaea. The tree was calculated with nearly full-length sequences (>1,300 bp) by maximum-likelihood analysis in combination with filters excluding highly variable positions. Data in colored boxes give information about the distribution and abundance of sequence retrieval and (b) ANME gene sequences coding for the alpha subunit of methyl coenzyme M reductase (mcrA) to selected sequences of the domain archaea. The tree was generated from deduced amino acid sequences (>247 amino acids) by neighbor-joining analysis with a 30% amino acid frequency filter. mcrA tree by courtesy of Lo¨sekann. Bar, 10% estimated sequence divergence.

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ANAEROBIC OXIDATION OF METHANE WITH SULFATE

Anaerobic Oxidation of Methane with Sulfate, Figure 2 Epifluorescence micrographs of different ANME single cells and aggregates visualized by fluorescence in situ hybridization (FISH) or CARD-FISH. (a) Single ANME-1 cells living in a microbial mat from the Black Sea; (b) mat-type consortium formed by ANME-1 (red) and DSS cells (green); (c) aggregated ANME-1 (green) and ANME-2 cells (blue) in a microbial mat from the Black Sea; (d) single ANME-2a cells in an enrichment culture from Hydrate Ridge sediments; (e–k) mixed-type consortia of ANME-2a (red) and DSS (green) cells observed in different seep sediments; (l) single ANME-2a cells; (m) monospecific ANME-2a aggregate; (n) corresponding DAPI staining in an enrichment culture from Hydrate Ridge sediments; (o–x) shell-type consortia of ANME-2c (red) and DSS (green) cells of different sizes and structure observed in different seep sediments; (y–aa): ANME-3; (y) monospecific aggregation of ANME-3 at Haakon Mosby mud volcano, (z, aa) ANME-3/Desulfobulbus consortia; and (ab): DAPI staining of a large ANME-2c aggregate. Unless otherwise indicated, scale bar 10 m.

ANAEROBIC OXIDATION OF METHANE WITH SULFATE

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Anaerobic Oxidation of Methane with Sulfate, Figure 3 Phylogenetic tree showing the affiliations of ANME-sulfate-reducing bacteria partner 16S rRNA gene sequences to selected reference sequences of the Deltaproteobacteria. The tree was calculated with nearly full-length sequences (>1,450 bp) by maximum-likelihood analysis in combination with filters excluding highly variable positions. Groups known as ANME partners are printed in boldface. Scale bar gives 10% estimated sequence divergence.

(Pernthaler et al., 2008). Lösekann et al. found evidence for ANME-3 aggregates with unidentified partners showing a mixed-type morphology in sediments from the Arctic mud volcano Haakon Mosby (Lösekann et al., 2007).

Habitats of ANME Since their discovery, the distribution of the ANME organisms has been studied intensively, mainly based on 16S rRNA gene phylogeny. More than 1800 16S rRNA gene sequences are available now from more than 50 different marine methane seeps (Hinrichs et al., 1999; Orphan et al., 2001a; Aloisi et al., 2002; Tourova et al., 2002; Mills et al., 2003, 2004, 2005; Inagaki et al., 2004; Heijs et al., 2005, 2007; Knittel et al., 2005; Lanoil et al., 2005; Stadnitskaia et al., 2005; Arakawa et al., 2006a, b; Fang et al., 2006; Lloyd et al., 2006; Martinez et al., 2006; Niemann et al., 2006a; Nunoura et al., 2006; Reed et al., 2006; Lösekann et al., 2007), vents (Takai and Horikoshi, 1999; Teske et al., 2002; Kelley et al., 2005; Brazelton et al., 2006; Inagaki et al., 2006a), and “sulfate–methane

transition zone” (SMTZ) (Thomsen et al., 2001; Niemann et al., 2005; Parkes et al., 2007) differing in e.g., temperature, methane flux, salinity, and pH. Furthermore, the presence of ANME organisms in the water column (Black Sea; Vetriani et al., 2003; Schubert et al., 2006a) and in a terrestrial mud volcano in the Carpathian mountains (Romania; Alain et al., 2006) has also been reported. A comparison of these reports shows that globally, the two most abundant and diverse phylogenetic groups of methane-oxidizing archaea are the ANME-1 and ANME-2 clades. The distribution of ANME-2a and ANME-2c is as wide as for ANME-1, but only few sequences have been reported belonging to subgroup ANME-2b. ANME-2b, originally defined on the basis of few sequences as a separate group (Orphan et al., 2001a), now seem to be rather a subgroup of ANME-2a than a monophyletic group (Figure 1a). Today it is known that ANME are present wherever methane and sulfate co-occur, at a wide range of environmental conditions (Knittel and Boetius, 2009). However,

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usually only one specific ANME group dominates a habitat and seems to be responsible for most of the measured AOM (Knittel et al., 2005). High rates of methane oxidation, sulfate reduction, and/or sulfide production can be used as an indicator for high densities of ANME populations (Boetius et al., 2008). Continuous high fluxes of both AOM substrates methane and sulfate at mM concentrations sustain high cell densities of >1010 ANME cells cm3 in the environment and can even form some of the densest and largest cell accumulations known to exist in nature (Michaelis et al., 2002).

Seep ecosystems Cold seeps form where tectonic or gravitational forces advect free gas, methane-rich porewater and/or mud upward into the sulfate-penetrated surface sediments and sometimes into the hydrosphere (Judd and Hovland, 2007; see Cold Seeps). The increased availability of dissolved methane at higher hydrostatic pressures increases the energy yield of AOM and leads to a natural enrichment of ANME populations. The products of AOM, sulfide and carbonate accumulate in the seabed, forming the typical features of cold seep ecosystems such as carbonate precipitates and high sulfide fluxes. The sulfide is oxidized chemically and microbially with oxygen, nitrate, or iron close to the seafloor surface. One of the most striking visual features of submarine cold seep ecosystems are mats of thiotrophic bacteria covering the seafloor. These mats are usually associated with high AOM rates and dense ANME communities inhabiting the underlying sediments (Sahling et al., 2002; Treude et al., 2003; Joye et al., 2004; Niemann et al., 2006b). Also, a variety of thiotrophic microbe–animal symbioses profit from AOM, such as siboglinid tubeworms and mytilid as well as vesicomyid bivalves (Sibuet and Olu, 1998). Thiotrophic mats and invertebrate symbioses may enhance the growth of ANME populations by rapidly removing the toxic endproduct sulfide, and by replenishing sulfate in the sediments (Treude et al., 2003; Cordes et al., 2005; Niemann et al., 2006b). In active cold seep sediments, AOM rates of 500–5,000 nmol CH4 cm3 day1 (>2 mol m2 year1) are reached, associated with very dense ANME populations of >1010 cells cm3 (Boetius et al., 2000; Knittel et al., 2005; Niemann et al., 2006b). Cold seep ecosystems from Eel River Basin (Hinrichs et al., 1999; Orphan et al., 2001b), Hydrate Ridge (Boetius et al., 2000; Knittel et al., 2003, 2005), Black Sea (Thiel et al., 2001, 2003, 2007; Michaelis et al., 2002; Blumenberg et al., 2005; Reitner et al., 2005a, b; Treude et al., 2005a, 2007), Gulf of Mexico (Orcutt et al., 2005; Lloyd et al., 2006; Martinez et al., 2006), the Tommeliten and Gullfaks area in the North Sea (Wegener et al., 2008), and mud volcanoes from the Mediterranean Sea (Omoregie et al., 2008) and the Barents Sea (Niemann et al., 2006b; Lösekann et al., 2007) have been intensively studied. With the exception of cold seeps in the Black Sea (see next) most of the studies indicate a

dominance of ANME-2 or ANME-3. A prominent example is the gas hydrate bearing sediment from the Hydrate Ridge. Hot spots of ANME-2 are surface layers just above the gas hydrates with up to 108 aggregates cm3. Interestingly, ANME-2 subgroups revealed preferences for either Beggiatoa (ANME-2a) or Calyptogena (ANME-2c) fields (Knittel et al., 2005) indicating that different environmental conditions select for different ANME groups. Two seep systems are known yet where other ANMEs dominate: (1) sediments overlaying a brine pool methane seep in the Gulf of Mexico are dominated by ANME-1b archaea. Here, the ANME-1b community was found in the sulfate-methane interface, where low methane concentrations of ca. 100–250 µM are present and coexist with sulfate concentrations of ca. 10 mM; and (2) microbial mats from Black Sea cold seeps. These seeps are unique ecosystems and AOM is important both in sediments and in the water column (Pimenov et al., 1997; Wakeham et al., 2003). Tall reef-like structures composed of porous carbonates and microbial mats are found on the seafloor. These mats have been shown to mediate AOM and consist mainly of densely aggregated ANME-1 cells and SRB (Michaelis et al., 2002; Blumenberg et al., 2004, 2005; Knittel et al., 2005; Reitner et al., 2005a, b; Treude et al., 2005a, 2007). Treude et al. (2007) combined radiotracer incubations, beta-microimaging, SIMS, and CARD-FISH to locate hot spots of methanotrophy. Incorporation of 14C from radiolabeled CH4 indicated a hot spot for methanotrophy close to the mat surface associated with a dominance of ANME-1 archaea (Treude et al., 2007). The mats, however, are very heterogeneous and also provide niches for ANME-2. Black nodules from the top of the reef seem to be dominated by ANME-2 as shown by specific 13C depleted lipids and FISH (Blumenberg et al., 2004). Reitner et al. found intracellular precipitation of iron sulfide (greigite) by bacteria growing in close association with ANME-2 and suggested iron cycling as an additional pathway involved in AOM (Reitner et al., 2005b). ANME-3 archaea are commonly found at submarine mud volcanoes (Niemann et al., 2006b; Heijs et al., 2007; Lösekann et al., 2007; Omoregie et al., 2008) but also at other cold seeps (Orphan et al., 2001a; Inagaki et al., 2004; Knittel et al., 2005), hydrothermal vents (Brazelton et al., 2006), and recently in subsurface sediments (Parkes et al., 2007).

Sulfate–methane transition zones (SMTZ) The most common ANME habitat on the Earth is the socalled SMTZ in the seabed (Thomsen et al., 2001; Ishii et al., 2004; Niemann et al., 2005, 2006a; Treude et al., 2005b; Parkes et al., 2007). SMTZ are found in all seabed horizons where the diffusive transport of methane from below and sulfate from above leads to a zone of AOM. Methane is completely consumed in the SMTZ, which may be found at decimeters to tens of meters below the seafloor (D'Hondt et al., 2004), depending on the depth

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of the methane production zone, on the transport velocity of methane and sulfate and on their consumption rates. In diffusive seabed systems, the distribution of ANME is restricted to SMTZ. Population densities of ANME are low with 70 elements dissolved in extant seawater, but only 6 ions make up >99% of all the dissolved salts (wt% in brackets): Na+ (30.51), Cl
(55.08), SO42
(7.69), Mg2+ (3.67), Ca2+ (1.17), K+ (1.10) (Hay et al., 2006). Introduction About 250 million years ago the continents on Earth were close together and formed Pangaea, a supercontinent, which persisted for about 100 million years and then fragmented. The land masses at that time were located predominantly in the southern hemisphere. The climate was arid and dry, and the average temperature is thought to have been several degrees higher than at present. This was one of the time periods in the history of the Earth, when

huge salt sediments formed. A total of about 1.3 million cubic kilometers of salt were estimated to have been deposited during the late Permian and early Triassic period alone (250–192 million years ago; Zharkov, 1981); newer research has discovered additional vast salt deposits, which were previously unknown – especially deposits below the Gulf of Mexico, and extensive Miocene salt (about 20 million years old) underlying the Mediterranean Sea, the Red Sea, and the Persian Gulf (Hay et al., 2006). The thickness of the ancient salt sediments can reach 1,000–2,000 m. When Pangaea broke up, land masses were drifting in latitudinal and northern direction. Mountain ranges such as the Alps, the Carpathians, and the Himalayas were pushed up by the forces of plate tectonics. In the Alpine basin and in the region of the Zechstein Sea, which covered northern Europe, no more salt sedimentation took place after the Triassic period. Dating of the salt deposits by sulfur-isotope analysis (ratio of 32S/34S as measured by mass spectrometry), in connection with information from stratigraphy, indicated a Permo–Triassic age for the Alpine and Zechstein deposits (Holser and Kaplan, 1966). This age was independently confirmed by the identification of pollen grains in the sediments (Klaus, 1974). Figure 1 shows pollen grains from extinct coniferous trees, which were found in Alpine Permian salt and also in Zechstein salt (Klaus, 1963).

Microorganisms and signature sequences from salt deposits As recently as 1981, Larsen (1981) described mined rock salts as free from bacteria, although isolations of halophilic microorganisms from ancient salt sediments had occasionally been reported since the early decades of the twentieth century (see Grant et al., 1998; McGenity et al., 2000). From Alpine Permian rock salt, which was collected from the salt mine in Bad Ischl, Austria, a haloarchaeon (see “Halobacteria – Halophiles”) was

Joachim Reitner & Volker Thiel (eds.), Encyclopedia of Geobiology, DOI 10.1007/978-1-4020-9212-1, # Springer Science+Business Media B.V. 2011

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DEEP BIOSPHERE OF SALT DEPOSITS

Deep Biosphere of Salt Deposits, Figure 1 Photomicrograph of winged pollen grains of the genus Lueckisporites, found in Alpine Permian salt deposits and in Zechstein salt. Upper panel: L. virkkiae, dimensions 60  40 mm; lower panel: L. microgranulatus, dimensions 64  35 mm (From Klaus, 1963 with permission.)

isolated, which was recognized as a novel species and named Halococcus salifodinae strain BIp (Denner et al., 1994). This was the first isolate from ancient rock salt, which was formally classified and deposited in international culture collections. Two independently isolated strains, Br3 (from solution-mined brine in Cheshire, England) and BG2/2 (from a bore core from the mine of Berchtesgaden, Germany) resembled Hc. salifodinae strain BIp in numerous properties, including the characteristic morphology of coccoid cells arranged in large clusters (see Figure 2, right panel, in “Halobacteria – Halophiles”); in addition, rock salt samples were obtained 8 years later from the same site and several halococci were recovered from these samples, which proved to be identical to strain BIp (Stan-Lotter et al., 1999). The data suggested that viable halophilic archaea, which belong to the same species, occur in geographically separated evaporites of similar geological age. Another halococcal isolate from the Bad Ischl salt formation, which differed from the previously described strains, was identified as a novel species and named Halococcus dombrowskii (Stan-Lotter et al., 2002). Hc. salifodinae and Hc. dombrowskii have so far not been found in any hypersaline surface waters, or any location other than salt mines. Recently, a non-coccoid novel haloarchaeon, Halobacterium noricense was obtained from a freshly

Deep Biosphere of Salt Deposits, Figure 2 Drilled bore cores, stored in a wooden box, from the Alpine salt mine in Altaussee, Austria obtained from about 500 m below surface. Pink portions represent halite (NaCl), containing traces of hematite; grayish portions contain mostly anhydrite (CaSO4).

drilled bore core (Figure 2) at the salt mine in Altaussee, Austria (Gruber et al., 2004). Other isolates from ancient salt deposits include a single rod-shaped Halobacterium from 97 000 year old rock salt in the USA (Mormile et al., 2003), which was deemed to resemble the wellcharacterized Halobacterium salinarum NRC-1 (see “Halobacteria – Halophiles”). From the Permian Salado formation in New Mexico, a novel strain, Halosimplex carlsbadense, was isolated (Vreeland et al., 2002). Although the microbial content of ancient rock salt is low – estimates range from 1 to 2 cells/kg of salt from a British mine (Norton et al., 1993) to 1.3  105 colony forming units (CFUs) per kg of Alpine rock salt (StanLotter et al., 2000), and up to 104 CFUs per g of Permian salt of the Salado formation (Vreeland et al., 1998), equivalent to a range of 1 pg to 10 mg of biomass per kg of saltthe reports showed that viable haloarchaeal isolates were obtained reproducibly by several groups around the world. The data support the hypothesis that the halophilic isolates from subterranean salt deposits could be the remnants of populations which inhabited once ancient hypersaline

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seas; in addition, they provide strong evidence against the notion that the recovered strains could be the result of laboratory contamination, since the isolates were obtained independently from different locations. The amplification of diagnostic molecules, such as the 16S rRNA genes of bacteria and archaea, by the polymerase chain reaction is now the standard method for obtaining material for subsequent nucleotide sequencing. For this technique it is not necessary to cultivate the microorganisms; the genes can be amplified by using DNA prepared from the material of interest. Analysis of dissolved Alpine rock salt with molecular methods was performed by extracting DNA and sequencing of 16S rRNA genes. The results provided evidence for the occurrence of numerous haloarchaea, which have not yet been cultured (Radax et al., 2001). Similarities of the 16S rDNA gene sequences were less than 90–95% to known sequences in about 37% of approximately 170 analyzed clones (Radax et al., 2001; Stan-Lotter et al., 2004); the remaining clone sequences were 98–99% similar to isolates from rock salts of various ages (McGenity et al., 2000) and to known haloarchaeal genera. In a similar experimental approach, using halite samples ranging in age from 11 to 425 millions of years, Fish et al. (2002) found haloarchaeal sequences and, in the older samples, also evidence for bacterial 16S rRNA genes which were related to the genera Aquabacterium, Leptothrix, Pseudomonas, and others. These data suggested the presence of a probably very diverse microbial community in ancient rock salt.

Long-term survival of cells The reports cited above provide evidence that viable extremely halophilic archaea were isolated from salt sediments, which are thought to have been deposited about tens of thousands or even millions of years ago. The fluid inclusions in Permian rock salt were reported to contain cations and anions in a similar composition as today’s seawater (Horita et al., 1991; Hay et al., 2006). While there is no direct proof that haloarchaea or other microorganisms have been entrapped in rock salt since its sedimentation, it would also be difficult to prove the opposite, namely that

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masses of diverse microorganisms entered the evaporites in recent times (see also McGenity et al., 2000, for further discussion). If a Permo–Triassic age is postulated for some of the haloarchaeal isolates and DNA signatures, then it becomes necessary to explain the biological mechanisms for such extreme longevity. Grant et al. (1998) discussed several possibilities, such as the formation of resting stages other than spores – since archaea are not known to form spores – or the maintenance of cellular functions with traces of carbon and energy sources within the salt sediments, which would imply an almost infinitely slow metabolism. At this time, there are no methods available to prove directly a great microbial age, whether it be a bacterium or a haloarchaeon. However, it can be shown, when simulating the formation of halite in the laboratory by drying salty solutions, which contained microorganisms that the cells accumulate within small fluid inclusions (Figure 3). The cells can be pre-stained with the fluorescent dyes of the LIVE/DEAD kit (Fendrihan et al., 2006), which provides information on the viability status of a cell (green fluorescence indicating viable cells); the procedure improves also the visualization of cells within crystals. The fluid inclusions were square or rectangular, as is common in the rectangular mineral halite, and the cells were rather densely packed within the fluid-filled spaces. From such experiments it appeared that the cells accumulated always in the fluid inclusions; there were no stained cells within the mineralic halite (Figure 3; Fendrihan et al., 2006). Suggestions have been made that fluid inclusions migrate within evaporites and thus, new nutrients might become accessible for the entrapped cells (McGenity et al., 2000).

Extraterrestrial halite and conclusion Traces of halite were found in the SCN meteorites (named after the locations where they were found – Shergotty in India, Nakhla in Egypt, and Chassigny in France), which stem from Mars (Treiman et al., 2000). The Monahans meteorite, which fell in Texas in 1998, contained macroscopic crystals of halite, in addition to potassium chloride and water inclusions (Zolensky et al., 1999). Recently,

Deep Biosphere of Salt Deposits, Figure 3 Localization of pre-stained haloarchaea in fluid inclusions of artificial halite. Cells were stained with the LIVE/DEAD BacLight kit prior to embedding. Low (left panel) and high (right panel) magnification of Halobacterium salinarum NRC-1 cells. Cells were observed with a Zeiss Axioskope fluorescence microscope.

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evidence for salt pools on the Martian surface was obtained (Osterloo et al., 2008). These results are intriguing – they suggest that the formation of halite with liquid inclusions could date back billions of years and occurred probably early in the formation of the solar system (Whitby et al., 2000). Could halophilic life have originated in outer space and perhaps traveled with meteorites, could haloarchaea have persisted in environments as they are found today on Mars? Viable extremely halophilic archaea, representing novel strains, were isolated repeatedly from Permo–Triassic and other ancient salt sediments, suggesting their capacity for long-term survival under dry conditions. Together with the discovery of extraterrestrial halite it appears thus feasible to include into the search for life on other planets or moons specifically a search for halophilic microorganisms.

Bibliography Denner, E. B. M., McGenity, T. J., Busse, H. J., Wanner, G., Grant, W. D., and Stan-Lotter, H., 1994. Halococcus salifodinae sp.nov., an archaeal isolate from an Austrian salt mine. International Journal of Systematic Bacteriology, 44, 774–780. Fendrihan, S., Legat, A., Gruber, C., Pfaffenhuemer, M., Weidler, G., Gerbl, F., and Stan-Lotter, H., 2006. Extremely halophilic archaea and the issue of long term microbial survival. Reviews in Environmental Science and Bio/technology, 5, 1569–1605. Fish, S. A., Shepherd, T. J., McGenity, T. J., and Grant, W. D., 2002. Recovery of 16S ribosomal RNA gene fragments from ancient halite. Nature, 417, 432–436. Erratum in: 2002. Nature, 420, 202. Gooding, J. L., 1992. Soil mineralogy and chemistry on Mars: possible clues from salts and clays in SNC meteorites. Icarus, 99, 28–41. Grant, W. D., Gemmell, R. T. and McGenity, T. J., 1998. Halobacteria: the evidence for longevity. Extremophiles, 2, 279–287. Gruber, C., Legat, A., Pfaffenhuemer, M., Radax, C., Weidler, G., Busse, H. J., and Stan-Lotter, H., 2004. Halobacterium noricense sp. nov., an archaeal isolate from a bore core of an alpine PermoTriassic salt deposit, classification of Halobacterium sp. NRC-1 as a strain of Halobacterium salinarum and emended description of Halobacterium salinarum. Extremophiles, 8, 431–439. Hay, W. W., Migdisov, A., Balukhovsky, A. N., Wold, C. N., Flögel, S., and Söding, E., 2006. Evaporites and the salinity of the ocean during the Phanerozoic: implications for climate, ocean circulation and life. Palaeogeography, Palaeoclimatology, Palaeoecology, 240, 3–46. Holser, W. T., and Kaplan, I. R., 1966. Isotope geochemistry of sedimentary sulfates. Chemical Geology, 1, 93–135. Horita, J., Friedman, T. J., Lazar, B., and Holland, H. D., 1991. The composition of Permian seawater. Geochimica et Cosmochimica Acta, 55, 417–432. Klaus, W., 1963. Sporen aus dem südalpinem Perm (Vergleichsstudie für die Gliederung nordalpiner Salzserien). Jahrbuch der Geologischen Bundesanstalt Wien, Band, 106, 229–361. Klaus, W., 1974. Neue Beiträge zur Datierung von Evaporiten des Oberperm. Carinthia II, 164, Jahrgang 84: 79–85. Larsen, H., 1981. The family Halobacteriaceae. In Starr, M. P., Stolp, H., Trüper, H. G., Balows A., and Schlegel H. G. (eds.). The Prokaryotes. A Handbook on Habitat, Isolation and Identification of Bacteria. Berlin: Springer, vol. I, pp. 985–994. McGenity, T. J., Gemmell, R. T., Grant, W. D., and Stan-Lotter, H., 2000. Origins of halophilic microorganisms in ancient salt deposits (MiniReview). Environmental Microbiology, 2, 243–250.

Mormile, M. R., Biesen, M. A., Gutierrez, M. C., Ventosa, A., Pavlovich, J. B., Onstott, T. C., and Fredrickson, J. K., 2003. Isolation of Halobacterium salinarum retrieved directly from halite brine inclusions. Environmental Microbiology, 5, 1094–1102. Norton, C. F., McGenity, T. J., and Grant, W. D., 1993. Archaeal halophiles (halobacteria) from two British salt mines. Journal of General Microbiology, 139, 1077–1081. Osterloo, M. M., Hamilton, V. E., Bandfield, J. L., Glotch, T. D., Baldridge, A. M., Christensen, P. R., Tornabene, L. L., and Anderson, F. S., 2008. Chloride-bearing materials in the southern highlands of Mars. Science, 319, 1651–1654. Radax, C., Gruber, C., and Stan-Lotter, H., 2001. Novel haloarchaeal 16S rRNA gene sequences from Alpine PermoTriassic rock salt. Extremophiles, 5, 221–228. Stan-Lotter, H., McGenity, T. J., Legat, A., Denner, E. B. M., Glaser, K., Stetter, K. O., and Wanner, G., 1999. Very similar strains of Halococcus salifodinae are found in geographically separated Permo-Triassic salt deposits. Microbiology, 145, 3565–3574. Stan-Lotter, H., Radax, C., Gruber, C., McGenity, T. J., Legat, A., Wanner, G., and Denner, E. B. M., 2000. The distribution of viable microorganisms in Permo-Triassic rock salt. In Geertman, R. M. (ed.), SALT 2000. 8th World Salt Symposium, Amsterdam: Elsevier Science BV, vol. 2, pp. 921–926. Stan-Lotter, H., Pfaffenhuemer, M., Legat, A., Busse, H. J., Radax, C., and Gruber, C., 2002. Halococcus dombrowskii sp. nov., an archaeal isolate from a Permo-Triassic alpine salt deposit. International Journal of Systematic and Evolutionary Microbiology, 52, 1807–1814. Stan-Lotter, H., Radax, C., McGenity, T. J., Legat, A., Pfaffenhuemer, M., Wieland, H., Gruber, C., and Denner, E. B. M., 2004. From intraterrestrials to extraterrestrials – viable haloarchaea in ancient salt deposits. In Ventosa A. (ed.), Halophilic Microorganisms. Berlin: Springer, pp. 89–102. Treiman, A. H., Gleason, J. D., and Bogard, D. D., 2000. The SNC meteorites are from Mars. Planetary and Space Science, 48, 1213–1230. Vreeland, R. H., Piselli, A. F., Mc-Donnough, S., and Meyers, S. S., 1998. Distribution and diversity of halophilic bacteria in a subsurface salt formation. Extremophiles, 2, 321–331. Vreeland, R. H., Straight, S., Krammes, J., Dougherty, K., Rosenzweig, W. D., and Kamekura, M., 2002. Halosimplex carlsbadense gen. nov., sp. nov., a unique halophilic archaeon, with three 16S rRNA genes, that grows only in defined medium with glycerol and acetate or pyruvate. Extremophiles, 6, 445–452. Whitby, J., Burgess, R., Turner, G., Gilmour, J. and Bridges, J., 2000. Extinct 129I halite from a primitive meteorite: evidence for evaporite formation in the early solar system. Science, 288, 1819–1821. Whitman, W. B., Coleman, D. C., and Wiebe, W. J., 1998. Prokaryotes: the unseen majority. Proceedings of the National Academy of Sciences of the United States of America, 95, 6578–6583. Zharkov, M. A., 1981. History of Paleozoic Salt Accumulation. Berlin: Springer. Zolensky, M. E., Bodnar, R. J., Gibson, E. K., Nyquist, L. E., Reese, Y., Shih, C. Y., and Wiesman, H., 1999. Asteroidal water within fluid inclusion-bearing halite in an H5 chondrite, Monahans (1998). Science, 285, 1377–1379.

Cross-references Archaea Astrobiology Bacteria Deep Biosphere of the Oceanic Deep Sea Extreme Environments

DEEP BIOSPHERE OF THE OCEANIC DEEP SEA

Halobacteria – Halophiles Saline Lakes Salinity History of the Earth’s Ocean Terrestrial Deep Biosphere

DEEP BIOSPHERE OF SEDIMENTS See entries “Terrestrial Deep Biosphere,” “Deep Biosphere of Salt Deposits,” “Deep Biosphere of the Oceanic Deep Sea,” and “Basalt (Glass, Endoliths).”

DEEP BIOSPHERE OF THE OCEANIC DEEP SEA Kristina Rathsack, Nadia-Valérie Quéric, Joachim Reitner University of Göttingen, Göttingen, Germany

Definition and overview Although used in many different ways, the term “biosphere” is principally defined either as zone in which life occurs, thereby overlapping the atmosphere, the hydrosphere, and the lithosphere, or as the entity of living organisms on Planet Earth. Both perceptions commonly focus on the Earth’s near-surface environment, with all domains sharing solar energy used in the process of photosynthesis. The deep-sea realm takes a special position in this context, as deep-sea pelagic and the majority of benthic organisms live in the ocean’s aphotic zone and inhabit the widespread abyssal plains, respectively. For a long time, their main food source has been considered to be based on particulate organic matter (POM) from the ocean’s surface primary production and its sedimentation to abyssal depths (Gage and Tyler, 1991 and references therein, D’Hondt et al., 2002, 2004). With the discovery of “ocean vents” in the late 1970s (Corliss et al., 1979), this general perspective was broadened by the perception of the enormous potential of chemical energy through the reaction of seawater, rock material, and fluids rising from the Earth’s interior. According to this concept of energy for life, the term ‘surface biosphere’ has been opposed to ‘subsurface biosphere’ (also commonly found in literature as ‘deep biosphere’). Following this definition, the deepseafloor with its highly diverse topography from heterotrophic to pure chemotrophic habitats has to be treated as a transition zone between both biospheres. Opposed to the “deep hot biosphere” (Gold, 1992), occurring by definition in oceanic as well as terrestrial subsurface environments, stands the “deep cold biosphere” as defined for permafrost sediments (Vorobyova et al., 1997) and ice cores from the depths of Lake Vostoc (Venter, 2001). Life in the deep sea Comprising approximately 65% of the Earth’s surface, the deep-sea environment is characterized by hyperbaric, aphotic, and low-temperature conditions and highly

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diverse seascapes. Canyons, seamounts, ridges, fractures, and trenches, but also biogeochemical oases such as cold seeps, mud volcanoes, carbonate mounds, brine pools, gas hydrates, hot vent systems, and deep-water coral reefs provide ample niches for a highly diverse pelagic and benthic deep-sea community (Tyler, 2003). It was only during the construction of the transoceanic telegraphic communication network that people realized the ocean’s topographic alterations and astonishing depths. In 1861, the repair of an overgrown cable from 1,800 m water depth in the Mediterranean finally aroused the scientific community which by then adhered to Edward Forbes’ theory on a completely ‘azotic’ zone below a water depth of 550 m. Though, 11 years had elapsed before the first global, scientific expedition onboard the “Challenger” (1872–1876) finally convinced people that a flourishing life in fact exists in the deep-sea realm. Numerous, further expeditions and a rushing development of technical facilities allowed deep-sea researches in 1960 to reach even the ocean’s deepest surveyed point, the Challenger Deep at 10.911 meters below sea level (mbsl), located at the southern end of the Mariana Trench within the western Pacific Ocean (Piccard and Dietz, 1961). Since then, several studies on large-scale patterns and the zoogeographical origins of deep-sea organisms evidenced a high macrobenthic diversity (Gage and Tyler, 1991 and references therein). These organisms display a depthdependent zonation as a result of basin age, deep currents (as barriers or dispersal), topographic boundaries, disturbance processes, and sedimentation in connection with depth-related environmental patterns (for review see Levin et al., 2001; Stuart et al., 2003). Macro- and meiofauna are loosing importance with increasing water and sediment depth, whereas microorganisms like bacteria, archaea, and fungi account for up to 90% of the deep-sea benthic biomass (Pfannkuche, 1992). Sinking particles may carry large numbers of microorganisms from upper zones (108–1010 cells m
2 d
1), inoculating deep marine surface sediments with an autotrophic and heterotrophic microbial community, as demonstrated by results from sediment traps (Turley and Mackie, 1995; Danovaro et al., 2000; Vanucci et al., 2001) or the deepseafloor (Lochte and Turley, 1988).

Particulate organic matter (POM) Due to the fact that most deep-sea benthic species are deposit feeders (Sanders and Hessler, 1969), the locally qualitatively and quantitatively, variable import of POM from the ocean’s surface waters plays a crucial role for macro-, meio-, and microorganisms living in deep surface sediments (Gooday and Turley, 1990). Mainly consisting of phytoplankton, marine snow, fecal pellets, (dead) zooplankton and molts, this material undergoes different steps of degradation during its passage from the photic, epipelagic (0–200 mbsl), through the mesopelagic (200– 1,000 mbsl) to the actual deep-sea zones, in particular the bathypelagic (1,000–4,000 mbsl), the abyssal

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(4,000–6,000 mbsl), and the hadal zone (6,000–11,000 mbsl). Depending on the residence time in the water column, the bioavailable part of POM finally reaching the deep-seafloor may be small (De La Rocha and Passow, 2007 and references therein). The refractory remainders such as animal skeletons are continuously accumulating at the seafloor and turn into deeply buried sediment over time, thereby representing the largest global reservoir organic carbon (Parkes et al., 2000 and references therein).

Deep-sea sediment types Grain size (Gray, 1974) and sediment heterogeneity (Etter and Grassle, 1992) may additionally govern community composition and distribution of macro-, meio-, and microorganisms in deep-sea benthic environments. In relation to their basic sources, deep-sea sediments may be biogenic (POM from pelagic primary production, benthic in-situ production), lithogenous (terrestrial weathering and transport by wind and rivers), hydrogenous (precipitation from seawater or pore water), volcanic, or cosmic (Seibold and Berger, 1996). According to grain size and settling velocity, lithogenous gravel and sandy fractions usually are deposited along the coast, while silt and clay are transported farther offshore through waves and currents, hence dominating the basically biogenous deep-sea sediments. Regional deviations may be linked to currents, downslope slides, submarine canyon dynamics, or to a release of ice-trapped rock material in polar waters (e.g., Ramseier et al., 2001). Covering almost one-half of the shelves and more than half of the deep ocean bottom, biogenous sediments mainly consist of calcite, aragonite, opal, and calcium phosphate, originating from foraminifera, diatoms, and radiolarians (Hay et al., 1988). Deep biosphere of deep-sea sediments Microbiological studies on sediment cores collected during several cruises of the Deep Sea Drilling Project (DSDP), the Ocean Drilling Program (ODP), and the Integrated Ocean Drilling Program (IODP) gave evidence for the presence of complex microbial communities in deeply buried marine sediments down to several hundred meters below seafloor (e.g., Whelan et al., 1986; Parkes et al., 1994; Roussel et al., 2008). Most striking, new insights into subsurface microbiology were gained during the ODP cruise Leg 201 to the equatorial Pacific Ocean and the continental margin of Peru, including sites recognized as most typical for oceanic subsurface environments (D’Hondt et al., 2002). A large fraction of the sub-seafloor bacteria has been proven to be alive and culturable, displaying turnover rates (based on sulfate reduction as dominating mineral process at these sites) comparable to surface sediment communities (D’Hondt et al., 2004; Schippers et al., 2005). After a logarithmic decline within the uppermost 6 meters below seafloor (mbsf) (Parkes et al., 1994) to about 40 mbsf (Schippers et al., 2005), bacterial cells have proven to be more or less evenly distributed down to

several hundred mbsf. Local peaks within these deeply buried sediments seem to mirror sulfate (diffusing from crustal fluids) and methane (from in-situ production) concentration shifts (Engelen et al., 2008). However, published variations of absolute cell numbers (by a factor of up to 3) have to be treated with caution: varying estimations not only depend on the geochemical conditions at the respective sampling sites, but also on the enumeration techniques applied. Calculations based on early results revealed that sub-seafloor sediments comprise – at least – half of all prokaryotic cells and up to one-third of the living biomass on Earth pointing to a slow-growing strategy of high biomass in areas of low-energy flux (Whitman et al., 1998). The prokaryotic community in deeply buried sediments can not exclusively be traced back to contaminations from biologically active surface layers or reactivation of spores and dormant cells (Parkes et al., 2000 and references therein). Porewater chemistry data obtained from sites throughout the world’s oceans (ODP, DSDP) showed that sulfate reduction, methanogenesis, and fermentation are the principal degradative metabolic processes in subsurface sediments. These results give evidence for significant lower metabolic rates for the subsurface compared to the surface biosphere and for methanogenesis becoming more important the more sulfate gets depleted with increasing sediment depth (D’Hondt et al., 2002 and references therein).

Windows to the subsurface biosphere Hydrothermal vents The discovery of the “ocean vents” near Galapagos Island (Corliss et al., 1979) was the first proof for the active movement of the gigantic oceanic plates of the Earth’s crust creating series of cracks in the ocean floor, teeming with life. At these discharge areas, hydrothermal fluids with temperatures of more than 400 C (Haase et al., 2007) mix up with the cold ocean seawater, resulting in a precipitation of dissolved metals and in the formation of characteristic chimneys over time. Iron and sulfide precipitates turn the smokers black (“black smokers,” Figure 1), while barium, calcium, and silicon minerals result in “white smokers.” Thermal precipitation and/or direct magma degassing of H2, H2S, CH4, CO, and CO2 in combination with oxygen as electron acceptor provide enough energy to support a highly productive and physiologically diverse chemoautotrophic microbial community (Reysenbach and Shock, 2002). As the highly diverse and dense hot vent macrofauna (e.g., vestimentiferan tubeworms, bivalve mollusks, provannid gastropods, alvinellid polychaete, and bresiliid shrimps) cannot feed on the released chemicals themselves, they either feed on chemoautotrophic microbes or host them as symbionts. The predominant endosymbionts are mesophilic to moderately thermophilic chemoautotrophs (mostly Gammaproteobacteria), whereas most

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Cold seeps and mud volcanoes Just a few years after the discovery of the hydrothermal vent systems, cold seep ecosystems were reported from active and passive continental margins and subduction zones all over the world (Aharon, 1994 and references therein). High-pressure, low oxygen and low-temperature conditions favour the formation of marine gas hydrates. In the subsurface realm, such gas reservoirs are stored in a crystalline form, whereas they get dissolved in pore waters and finally leave the sediment surface in gaseous form. High fluxes of methane, sulfide, and other reduced elements characterize these ecosystems such as cold seeps, hydrocarbon vents and mud volcanoes, often leaving mineral precipitation in their immediate surroundings. Coupled to sulfate reduction, rich bacterial and archaeal communities perform anaerobic oxidation of hydrocarbons, but predominately of methane (Boetius et al., 2000; Borowski et al., 2000; Treude et al., 2005).

Conversion of methane is mainly mediated by two different groups of anaerobic methanotrophic archaea (ANME-I and ANME-II) (Nauhaus et al., 2005), forming syntrophic consortia with the sulfate-reducing bacteria (SRB) Desulfosarcina and Desulfococcus (Hinrichs et al., 1999; Boetius et al., 2000; Michaelis et al., 2002; Knittel et al., 2003). The methane-emitting Haakon Mosby Mud Volcano (HMMV, Barents Sea) has shown to harbor three key communities in methane conversion such as aerobic, methanotrophic bacteria (Methylococcales), anaerobic methanotrophic archaea (ANME-2) thriving below siboglinid tubeworms, and a previously undescribed clade of archaea (ANME-3) associated with bacterial mats (Niemann et al., 2006). Similarly, some cold seeps on the deeper Black Sea shelf are characterized by intense methane bubble discharge, mainly related to microbial methanogenesis (Pape et al., 2008 and references therein). Diffuse gas seeps in more shallow, oxic Black Sea waters often exhibit a netlike coverage of microbial mats similar to Beggiatoa-mats observed at HMMV (Figure 2). Beggiatoa spp. are discussed as keystone members of seep communities owing to their ability to (directly and indirectly) influence the metabolic activity of d-Proteobacteria, Planctomycetales, and ANME archaea by providing sulfate and ammonia as reactants (Mills et al., 2004). The question remains, to which extent such seep systems influence the global methane cycle, as the quantification of bubble dissolution and/or the release of methane-rich pore fluids from the sediment into the hydrosphere is difficult to achieve (Vogt et al., 1999; Reeburgh, 2007). Niemann et al. (2006) estimated that methanotrophy at active marine mud volcanoes consumes less than 40% of the total methane flux, due to limitations

Deep Biosphere of the Oceanic Deep Sea, Figure 1 Black smoker “Candelabra,” Logachev hydrothermal field, Mid-Atlantic Ridge (3,000 m water depth). (By courtesy of MARUM, Center for Marine Environmental Sciences, Bremen, Germany.)

Deep Biosphere of the Oceanic Deep Sea, Figure 2 Shallowwater cold seep area, Ukrainian shelf, Black Sea. White, net-like microbial mats are constructed of sulfide oxidizing bacteria (“Beggiatoa”). (By courtesy of Karin Hissmann and Ju¨rgen Schauer, Jago Team, Leibniz-Institut fu¨r Meereswissenschaften (IFM-GEOMAR) Kiel, Germany.)

episymbionts belong to the Epsilonproteobacteria, which can oxidize H2 and sulfur compounds while reducing oxygen, nitrate, and sulfur compounds (for review, see Nakagawa and Takai, 2008). The vent habitat proved to harbor methanogens (Methanococcus), sulfate-reducers (Archaeoglobus), and facultative autotrophs and heterotrophs such as the thermophilic aerobic Thermus and Bacillus (Harmsen et al., 1997) or, for example, Thermococcus, Phyrococcus, Desulfurococcus (Prieur et al., 1995; Teske et al., 2000; Nercessian et al., 2003; Schrenk et al., 2003). Generally detected archaeal phylotypes were affiliated with hyperthermophilic Crenarchaeota, Euryarchaeota Group I, II, III (Takai and Horikoshi, 1999) and the “Deep-sea Hydrothermal Vent Euryarchaeotal Group” (Hoek et al., 2003).

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of the relevant electron acceptors in the upward flowing, sulphate- and oxygen-free fluids.

Deep biosphere of the oceanic crust The fact that microorganisms are present in the subsurface realm had been reported decades ago in terrestrial subsurface environments (Farrell and Turner, 1931; Lipman, 1931). Early drilling operations performed for commercial purposes such as mining, oil and hot water recovery, and the search for underground waste repositories reported on the existence of a large community of microorganisms obviously involved in geochemical processes in the deep biosphere (Gold, 1992; Pedersen, 1993 and references therein). Hence, it was only in the early 1990s that scientists started to focus on the investigation of prospering life beneath the Earth’s crust, thanks to a chance encounter of a deep oceanic, volcanic eruption during a dive onboard the submersible Alvin, releasing white microbial bulk mats (Haymon et al., 1993). The upper layers of the oceanic crust are characterized by high basaltic porosity, hosting a vast hydrothermal reservoir (Johnson and Pruis, 2003) inhabited by a microbial community composed of species that are also found in deep-sea waters, sediments, and the deep oceanic crust (Thorseth et al., 2001; Huber et al., 2006). Among the most prominent anaerobic thermophiles indigenous for the oceanic crust, the Ammonifex group of bacteria (Nakagawa et al., 2006) or groups within Crenarchaeota, Euryarchaeota, and Korarchaeota (Ehrhardt et al., 2007). Since 3.5 billion years, basaltassociated glass textures and vesicular cavities within the basaltic matrix provide niches for microbial colonization (Furnes et al., 2004; Peckmann et al., 2008). For instance, the fossil record of the oceanic crust even gives evidence for a previous fungal life in deep ocean basaltic rocks (Schumann et al., 2004). Much effort has been put into the investigation of the deep biosphere of the deep sea during the past 20 years. However, we still are neither aware of the final composition of the living subsurface community, nor of its interrelationship to, for example, crustal fluid-derived compounds, nor of its global impact. Bibliography Aharon, P., 1994. Geology and biology of modern and ancient submarine hydrocarbon seeps and vents: an introduction. Geomarine Letters, 14(2), 69–73. Boetius, A., Ravenschlag, K., Schubert, C. J., Rickert, D., Widdel, F., Gieseke, A., Amann, R., Jorgensen, B. B., Witte, U., and Pfannkuche, O., 2000. A marine microbial consortium apparently mediating anaerobic oxidation of methane. Nature, 407(6804), 623–626. Borowski, W. S., Cagatay, N., Ternois, Y., and Paull, C. K., 2000. Data report: carbon isotopic composition of dissolved CO2, CO2 gas, and methane, Blake-Bahama Ridge and northeast Bermuda Rise, ODP Leg 172. Proceedings of the Ocean Drilling Program, Scientific Results 172. http://www.odp.tamu.edu/publications/172_SR/chap_03/chap_03.htm. Corliss, J. B., Dymond, J., Gordon, L. I., Edmond, J. M., von Herzen, R. P., Ballard, R. D., Green, K., Williams, D.,

Bainbridge, A., Crane, K., and VanAndel, T. H., 1979. Submarine thermal springs on the Galápagos rift. Science, 203, 1073–1083. Danovaro, R., Della Croce, N., Dell’Anno, A., Mauro F., Marrale, D., and Martorano, D., 2000. Seasonal changes and biochemical composition of the labile organic matter flux in the Cretan Sea. Progress in Oceanography, 46, 259–278. De La Rocha, C., and Passow, U., 2007. Factors influencing the sinking of POC and the efficiency of the biological carbon pump. Deep Sea Research II, 54, 639–658. D’Hondt, S., Rutherford, S., and Spivack, A. J., 2002. Metabolic activity of subsurface life in deep-sea sediments. Science, 295(5562), 2067–2070. D’Hondt, S., Jorgensen, B. B., Miller, D. J., Batzke, A., Blake, R., Cragg, B. A., Cypionka, H., Dickens, G. R., Ferdelman, T., Hinrichs, K.-U., Holm, N. G., Mitterer, R., Spivack, A., Wang, G., Bekins, B., Engelen, B., Ford, K., Gettemy, G., Rutherford, S. D., Sass, H., Skilbeck, C. G., Aiello, I. W., Guerin, G., House, C. H., Inagaki, F., Meister, P., Naehr, T., Niitsuma, S., Parkes, R. J., Schippers, A., Smith, D. C., Teske, A., Wiegel, J., Padilla, C. N., and Acosta, J. L. S., 2004. Distributions of microbial activities in deep subseafloor sediments. Science, 306(5705), 2216–2221. Engelen, B., Ziegelmüller, K., Wolf, L., Köpke, B., Gittel, A., Cypionka, H., Treude, T., Nakagawa, S., Inagaki, F., Lever, M. A., and Steinsbu, B. O., 2008. Fluids from the oceanic crust support microbial activities within the deep biosphere. Geomicrobiology Journal, 25(1), 56–66. Ehrhardt, C. J., Haymon, R. M., Lamontagne, M. G., and Holden, P. A., 2007. Evidence for hydrothermal Archaea within the basaltic flanks of the East Pacific Rise. Environmental Microbiology, 9, 900–912. Etter, R. J., and Grassle, J. F., 1992. Patterns of species diversity in the deep sea as a function of sediment particle size diversity. Nature, 360, 576–578. Farrell, M. A., and Turner, H. G., 1931. Bacteria in anthracite coal. Journal of Bacteriology, 23, 155pp. Furnes, H., Banerjee, N. R., Muehlenbachs, K., Staudigel, H., and de Wit, M., 2004. Early life recorded in Archean pillow lavas. Science, 304, 578–581. Gage, J. D., and Tyler, P. A., 1991. Deep-sea Biology – A Natural History of Organisms at the Deep-Sea Floor. Cambridge: Cambridge University Press. Gold, T., 1992. The deep, hot biosphere. Proceedings of the National Academy of Sciences of the United States of America, 89(13), 6045–6049. Gooday, A. J., and Turley, C. M., 1990. Responses by benthic organisms to inputs of organic material to the ocean floor: a review. Philosophical Transactions of the Royal Society of London. Series A, Mathematical and Physical Sciences, 331(1616), 119–137. Gray, J. S., 1974. Animal–sediment relationships. Oceanography and Marine Biology: An Annual Review, 12, 223–261. Haase, K. M., Petersen, S., Koschinsky, A., Seifert, R., Devey, C. W., Keir, R., Lackschewitz, K. S., Melchert, B., Perner, M., Schmale, O., Süling, J., Dubilier, N., Zielinski, F., Fretzdorff, S., Garbe-Schönberg, D., Westernströer, U., German, C. R., Shank, T. M., Yoerger, D., Giere, O., Kuever, J., Marbler, H., Mawick, J., Mertens, C., Stöber, U., and Walter, M., 2007. Young volcanism and related hydrothermal activity at 5 S on the slow-spreading southern Mid-Atlantic ridge. Geochemistry Geophysics Geosystems, 8(11), 1–17. Harmsen, H. J. M., Prieur, D., and Jeanthoni, C., 1997. Distribution of microorganisms in deep-sea hydrothermal vent chimneys investigated by whole-cell hybridization and enrichment culture of thermophilic subpopulations. Applied and Environmental Microbiology, 63(7), 2876–2883.

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Hay, W. W., Sloan II, J. L., and Wold, C. N., 1988. Mass/age distribution and composition of sediments on the ocean floor and the global rate of sediment subduction. Journal of Geophysical research, 93(B12), 14.933–14.940. Haymon, R., Fornari, D., Von Damm, K., Lilley, M., Perfit, M., Edmond, J., Shanks III, W. C., Lutz, R., Grebmeier, J., Carbotte, S., Wright, D., McLaughlin, E., Smith, M., Beedle, N., and Olson, E., 1993. Volcanic eruption of the mid-ocean ridge along the East Pacific Rise at 9 45–520 Ν: direct submersible observation of seafloor phenomena associated with an eruption event in April, 1991. Earth and Planetary Science Letters, 119, 85–101. Hinrichs, K. U., Hayes, J. M., Sylva, S. P., Brewer, P. G., DeLong, E. F., 1999. Methane-consuming archaebacteria in marine sediments. Nature 398, 802–805. Hoek, J., Banta, A., Hubler, F., and Reysenbach, A. L., 2003. Microbial diversity of a sulphide spire located in the Edmond deep-sea hydrothermal vent field on the Central Indian Ridge. Geobiology, 1, 119–127. Huber, J. A., Johnson, H. P., Butterfield, D. A., and Baross, J. A., 2006. Microbial life in ridge flank crustal fluids. Environmental Microbiology, 8, 88–99. Johnson, H. P., and Pruis, M. J., 2003. Fluxes of fluid and heat from the oceanic crustal reservoir. Earth and Planetary Science Letters, 216, 565–574. Knittel, K., Boetius, A., Lemke, A., Eilers, H., Lochte, K., Pfannkuche, O., and Linke, K., 2003. Activity, distribution, and diversity of sulfate reducers and other bacteria in sediments above gas hydrate (Cascadia Margin, Oregon). Geomicrobiology Journal, 20, 269–294, Kulm et al., 1986. Levin, L. A., Etter, R. J., Rex, M. A., Gooday, A. J., Smith, C. R., Pineda, J., Stuart, C. T., Hessler, R. R., and Pawson, D. L., 2001. Environmental influences on regional deep-sea species diversity. Annual Review of Ecology and Systematics, 32, 51–93. Lipman, C. B., 1931. Living micro-organisms in ancient rocks. Journal of Bacteriology, 22, 183pp. Lochte, K., and Turley, C. M., 1988. Bacteria and cyanobacteria associated with phytodetritus in the deep sea. Nature, 333, 67–68. Michaelis, W., Seifert, R., Nauhaus, K., Treude, T., Thiel, V., Blumenberg, M., Knittel, K., Gieseke, A., Peterknecht, K., Pape, T., Boetius, A., Amann, R., Jorgensen, B. B., Widdel, F., Peckmann, J., Pimenov, N. V., and Gulin, M. B., 2002. Microbial reefs in the Black Sea fueled by anaerobic oxidation of methane. Science, 297(5583), 1013–1015. Mills, H. J., Martinez, R. J., Story, S., and Sobecky, P. A., 2004. Identification of members of the metabolically active microbial populations associated with Beggiatoa species mat communities from Gulf of Mexico cold-seep sediments. Applied and Environmental Microbiology, 70(9), 5447–5458. Nakagawa, S., and Takai, K., 2008. Deep-sea vent chemoautotrophs: diversity, biochemistry and ecological significance. FEMS Microbiology Ecology, 65(1), 1–14. Nakagawa, S., Inagaki, F., Suzuki, Y., Steinsbu, B. O., Lever, M. A., Takai, K., Engelen, B., Sako, Y., Wheat, C. G., and Horikoshi, K., 2006. Microbial community in black rust exposed to hot ridge flank crustal fluids. Applied and Environmental Microbiology, 72(10), 6789–6799. Nauhaus, K., Treude, T., Boetius, A., and Krüger, M., 2005. Environmental regulation of the anaerobic oxidation of methane: a comparison of ANME-I and ANME-II communities. Environmental Microbiology, 7(1): 98–106. Nercessian, O., Reysenbach, A. L., Prieur, D., and Jeanthon, C., 2003. Archaeal diversity associated with in situ samplers deployed on hydrothermal vents on the East Pacific Rise (13 ?N). Environmental Microbiology, 5(6), 492–502.

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Niemann, H., Lösekann, T., de Beer, D., Elvert, M., Nadalig, T., Knittel, K., Amann, R., Sauter, E. J., Schlüter, M., Klages, M., Foucher, J. P., and Boetius, A., 2006. Novel microbial communities of the Haakon Mosby mud volcano and their role as a methane sink. Nature, 443, 854–858. Pape, T., Blumenberg, M., Seifert, R., Bohrmann, G., and Michaelis, W., 2008. Marine methane biogeochemistry of the Black Sea: a review. In: Dilek, Y., Furnes, H., and Muehlenbachs, K. (eds.), Links Between Geological Processes, Microbial Activities & Evolution of Life. Berlin: Springer, pp. 281–311. Parkes, R. J., Cragg, B. A., Bale, S. J., Getlifff, J. M., Goodman, K., Rochelle, P. A., Fry, J. C., Weightman, A. J., and Harvey, S. M., 1994. Deep bacterial biosphere in Pacific Ocean sediments. Nature, 371(6496), 410–413. Parkes, J. R., Cragg, B. A., and Wellsbury, P., 2000. Recent studies on bacterial populations and processes in subseafloor sediments: a review. Hydrogeology Journal, 8, 11–28. Peckmann, J., Bach, W., Behrens, K., and Reitner, J., 2008. Putative cryptoendolithic life in Devonian pillow basalt, Rheinisches Schiefergebirge, Germany. Geobiology, 6, 125–135. Pedersen, K., 1993. The deep subterranean biosphere. EarthScience Reviews, 34(4), 243–260. Pfannkuche, O., 1992. Organic carbon flux through the benthic community in the temperate abyssal northeast Atlantic. In: Rowe, G. T., and Pariente, V. (eds.), Deep-Sea Food Chains and the Global Carbon Cycle. The Netherlands: Kluwer, pp. 183–198. Piccard, J., and Dietz, R. S., 1961. Seven Miles Down. New York: Putnam. Prieur, D., Erauso, G., and Jeanthon, C., 1995. Hyperthermophilic life at deep-sea hydrothermal vents. Planetary and Space Science, 43(1–2), 115–122. Ramseier, R. O., Garrity, C., and Martin, T., 2001. An overview of sea-ice conditions in the Greenland Sea and the relationship of oceanic sedimentation to the ice regime. In: Schäfer, P., Ritzrau, W., Schlüter, M., and Thiede, J. (eds.), The Northern North Atlantic – A Changing Environment. Berlin: Springer, pp. 19–38. Reeburgh, W. S., 2007. Oceanic methane biogeochemistry. Chemical Reviews, 107(2), 486–513. Reysenbach, A. L., and Shock, E., 2002. merging genomes with geochemistry in hydrothermal ecosystems. Science, 296(5570), 1077–1082. Roussel, E. G., Cambon-Bonavita, M. A., Querellou, J., Cragg, B. A., Webster, G., Prieur, D., and Parkes, R. J., 2008. Extending the sub–sea-floor biosphere. Science, 320(5879), 1046. Sanders, H. L., and Hessler, R. R., 1969. Ecology of the deep-sea benthos. Science, 163(3874), 1419–1424. Schippers, A., Neretin, L. N., Kallmeyer, J., Ferdelman, T. G., Cragg, B. A., Parkes, R. J., and Jørgensen, B. B., 2005. Prokaryotic cells of the deep sub-seafloor biosphere identified as living bacteria. Nature, 433(7028), 861–864. Schrenk, M. O., Kelley, D. S., Delaney, J. R., and Baross, J. A., 2003. Incidence and diversity of microorganisms within the walls of an active deep-sea sulfide chimney. Applied and Environmental Microbiology, 69, 3580–3592. Schumann, G., Manz, W., Reitner, J., and Lustrino, M., 2004. Ancient fungal life in North Pacific Eocene Oceanic Crust. Geomicrobiology Journal, 21, 241–246. Seibold, E., and Berger, W. H., 1996. The Sea Floor – An Introduction to Marine Geology, 3rd edn. Berlin/Heidelberg/New York: Springer. Stuart, C. T., Rex, M. A., and Etter, R. J., 2003. Large-scale spatial and temporal patterns of deep-sea benthic species diversity. In: Tyler, P. A. (ed.), Ecosystems of the Deep Oceans. Amsterdam: Elsevier, pp. 295–311.

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Cross-references Anaerobic Oxidation of Methane with Sulfate Archaea Bacteria Basalt (Glass, Endoliths) Beggiatoa Biofilms Carbon Cycle Carbon (Organic, Cycling) Chemolithotrophy Cold Seeps Deep Biosphere of Salt Deposits Extreme Environments Hydrothermal Environments, Marine Methane Oxidation (Aerobic) Microbial Biomineralization Microbial Degradation Microbial Mats Microbialites, Modern Microbialites, Stromatolites, and Thrombolites Mud Mounds

Piezophilic Bacteria Sulfide Mineral Oxidation Sulfur Cycle Terrestrial Deep Biosphere

DEEP FLUIDS See entries “Terrestrial Deep Biosphere,” “Deep Biosphere of Salt Deposits,” and “Deep Biosphere of the Oceanic Deep Sea.”

DEGRADATION (OF ORGANIC MATTER) Transformations of organic matter within the range of temperatures, pressures and environmental conditions found at or near earth surface environments. If biological activity is confirmed to cause the transformations, termed biodegradation. See entries “Carbon (Organic, Cycling)" and “Carbon (Organic, Degradation)”, and “Microbial degradation (of organic matter)” for further reading.

DENITRIFICATION Denitrification is the biological reduction of nitrate (NO3
) to N2 and, to a minor extent, other gaseous species such as N2O. See entry “Nitrogen” for further reading.

DESERT VARNISH Randall S. Perry Imperial College, London, UK

Synonyms Rock Varnish, Rock Glaze, Silica Glaze, and Schutzrinde were introduced in 1891, from a translation of the French term manteau protecteur. “Schutzrinde” for all dark coatings, including “Wustenlack” a thin, polished coating that may be an eolian polished patina; “Dunkle Rinden” as a dark brown to black coating, possibly more similar to Sonoran and Mojave desert varnish; and “Schweinfurth,” which describes black coatings on rocks in Egypt that have spread on surrounding sands. Definition Desert Varnish. In arid environments such as Death Valley (California), rocks are covered with black opalescent desert varnish (Figure 1). Desert varnishes have been found in all continents, in locations such as the Gobi, Sonoran, Mojave, Namibian, Victorian, and Atacama

DESERT VARNISH

Desert Varnish, Figure 1 California desert varnish.

Deserts. The dark, lustrous coatings have attracted the interest of scientists for centuries. The German naturalist and explorer, Alexander Humboldt, observed desert varnish on a transatlantic expedition and questioned how this enigmatic feature would have formed. His contemporary, Darwin (1887), also engaged in the search for explanations for this unusual rock coating. To date, many other noteworthy scientists have examined desert varnish and have commented on its bulk chemistry, the arid conditions in which it forms and the concentration of manganese that makes it opaque and causes it to be black (cf. Perry et al., 2006, 2007; Perry, 1979; Staley et al., 1992). Geochemists, planetary geologists, and microbiologists as well as archeologists are interested in desert varnish, as petroglyphs are incised in varnish coatings all over the world. Yet, despite years of scientific effort, the origin of desert varnish is still shrouded in controversy. Most investigators have looked into biological causes where microbes create varnish and preferentially concentrate manganese relative to the local soils and rocks. However, more modern theories propose a nonbiological origin with sequential episodes of inorganic chemical reactions dominating formation processes. “Desert varnish” is used here to define coatings that appear dark (reddish brown, chocolate, black, and blueblack). Reasons for not abandoning desert varnish as a term are both because of its historical use and because

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it seems a most apt description for dark surfaces formed in subaerial conditions in hot and cold, and high and low elevation deserts. It must be emphatically stated that this definition of darkness is a macrovisual phenomenon. Close inspection of varnish coatings exposes the heterogeneity of the surface composition. Desert varnish is a thin sedimentary deposit (50 MPa) bacteria and archeaea (Fang and Bazylinski, 2008). Whereas, as a rule, the relative growth rate of these organisms decreases with pressure, the upper pressure limit of life has not yet been determined (Yayanos, 1998; Schrenk et al., 2010). High pressures affect biological systems by making the structures (e.g., membranes) more compact, as opposed to the effects of high temperature. Pressurization hinders any process resulting in a positive volume change, and vice versa. For example, if a reaction is accompanied by a volume decrease of 300 mL mol1, it is enhanced more than 200,000-fold by applying a pressure of 100 MPa (10,000 m water depth; Abe et al., 1999). Likewise, hydrostatic pressure has been shown to exert a considerable influence on many protein–protein interactions, the efficacy of enzymatic catalysis, replication, and translation. Piezophiles have therefore evolved specific adaptations, for example, in terms of membrane lipid composition and cell division. Pressure has also a significant effect on microbialmediated redox reactions, and metabolic versatility appears to be a specific adaptation to deep environments (Fang and Bazylinski, 2008; Lauro and Bartlett, 2008). For detailed reading, please refer to entry “Piezophilic Bacteria.” Extremely cold environments Environments with temperatures below 5 C for prolonged periods of time, such as cold polar regions, glaciers, seaice, deep-sea sediments, and permafrost soils are considered extremely cold environments. Organisms capable of growth and reproduction in these settings are called psychrophiles (cryophiles). They depend on low temperatures (< 0 C to 20 C) and have a growth optimum below 15 C (Morita, 1975). The lowest temperature limit for life seems to be around 20 C and has been reported for bacteria living in permafrost soil and in sea-ice (D’Amico

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et al., 2006). Psychrophiles are found in all three domains of life. Eukaryotes such as algae (e.g., sea-ice diatoms, the “snow alga” Chlamydomonas nivalis), protozoans, fungi, and even small metazoans frequently occur, but psychrophiles are most abundant among diverse lineages within the bacteria (e.g., Pseudomonas spp., Vibrio spp., several cyanobacterial genera) and, less often cited, archaea (e.g., Methanogenium and Methanococcus) (Garrison, 1991; D’Amico et al., 2006). Main coldinduced challenges posed to psychrophiles within their habitats encompass (i) reduced enzyme activity, (ii) decreased membrane fluidity, (iii) altered transport of nutrients and waste products, (iv) decreased rates of transcription, translation, and cell division, (v) protein colddenaturation and inappropriate protein folding, (viii) intracellular ice formation, and (ix) low water availability (see D’Amico et al., 2006, for a review). Psychrophilic organisms have successfully evolved strategies to overcome these negative effects of low temperatures. Such strategies involve modifications of the cell membrane composition toward a higher content of unsaturated, branched, or short-chain fatty acids, and large polar head groups (Chintalapati et al., 2004). Psychrophiles also synthesize specific antifreeze proteins, trehalose, and extracellular polymeric substances (EPS), which play an important role as cryoprotectants to keep the intercellular space liquid and protect the DNA at temperatures below water’s freezing point (see D’Amico et al., 2006, for a review). For further reading, please refer to “Permafrost Microbiology.”

Extremely dry environments Environments without free water are considered extremely dry environments and they include hot and cold deserts, and some terrestrial endolithic habitats (see also Chapter “Endoliths”). Organisms dependent on very dry environments are termed xerophiles, whereas those tolerating only temporary desiccation are referred to as “xerotolerant.” Most organisms thriving in very dry environments are actually xerotolerants, which rely on at least periodically available free water. A strategy to cope with prolonged periods of dryness is to enter the state of anhydrobiosis, which is characterized by little intracellular water and no metabolic activity. Organisms that can become anhydrobiotic are found among bacteria, yeast, fungi, plants, and even animals such as nematodes and the brine shrimp Artemia salina (Rothschild and Mancinelli, 2001). The terms “xerophile” and “xerotolerant” are often used to include organisms thriving under conditions of low water activity aw < 0.80). The aw is a measure of the amount of water within a medium that an organism can use to support growth. It represents the ratio of the water vapor pressure of the substrate to that of pure water under the same conditions and is expressed as a fraction (pure water, aw = 1; saturated NaCl, aw = 0.75). The lowest aw value recorded for growth to date was reported for the spoilage mould Xeromyces bisporus (aw = 0.61; see Grant, 2004 for a review). According to this

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definition, the most xerophilic organisms known, thrive in foods preserved by some form of dehydration or enhanced sugar levels, and in hypersaline environments where water availability is limited by a high concentration of salts (Grant, 2004). Whereas the former are dominated by xerophilic filamentous fungi and yeasts, high-salt environments are almost exclusively populated by prokaryotes. For the strategies employed by these organisms to cope with the osmotic stress exerted by these environments see above, and entry “Halobacteria – Halophiles.”

High-radiation environments Environments exposed to high doses of ionizing radiation are high-radiation environments. Ionizing radiation is radiation with sufficient energy to ionize molecules, most commonly ultraviolet (UV) radiation and natural radioactivity. When such radiation passes through (living) matter, ions and free radicals are produced that react rapidly and modify molecules (Cox and Battista, 2005). Ionizing radiation is therefore potentially detrimental for life, mainly due to DNA damage resulting from the generation of reactive oxygen species and the hydrolytic cleavage of water. Organisms that are capable of withstanding high doses of ionizing radiation are called radioresistant. There is no clear pattern of evolution among ionizing-radiation-resistant species, and they occur scattered over the three domains of life. Wellstudied microbial examples are the archaeon Thermococcus gammatolerans (Jolivet et al., 2003) and the bacterium Deinococcus radiodurans (Cox and Battista, 2005). Strategies to adapt to high doses of ionizing radiation involve (i) increasing the numbers of genome copies, (ii) tight spatial arrangement of nucleoids, (iii) accumulation of efficient radical scavengers, (iv) delaying DNA replication until damage repair has been completed, and (v) improved enzymatic genome-repair process. For further reading, please refer to entry “Radioactivity.” Summary “Extreme environment” refers to any setting that exhibits life conditions detrimental or fatal to higher organisms with respect to its physicochemical properties, in particular pH, temperature, pressure, saline concentrations, and radiation. Major classes of extreme environments encompass acidic (pH < 5), alkaline (pH > 9), hypersaline (salinity > 35%), pressurized (> 0.1 MPa), hot (> 40 C), cold ( +1,000 mV below pH 3) allows therefore only little energy generation (Kappler and Straub, 2005; Appelo and Postma, 2007). Nevertheless, representatives of different phyla of the Bacteria (e.g., Acidithiobacillus ferrooxidans, Sulfobacillus acidophilus) and the Archaea (Acidianus brierleyi, Sulfolobus metallicus) exploit Fe(II) in aerobic respiration. Moreover, three species of the genus Leptospirillum are obligate aerobic Fe(II)-oxidizing bacteria, i.e., Fe(II) is the only electron donor and molecular oxygen the only electron acceptor utilized (Johnson, 2007). All other acidophilic Fe(II)oxidizing prokaryotes make use of alternative electron donors (e.g., hydrogen, sulfur) and sulfur or Fe(III) as alternative electron acceptors. Species of acidophilic Fe(II)-oxidizing bacteria that also grow by dissimilatory reduction of Fe(III) were shown to catalyze the cycling of iron in laboratory experiments (Johnson et al., 1993).

Acidithiobacillus ferrooxidans was the first acidophilic aerobic Fe(II)-oxidizing bacterium that was isolated (Temple and Colmer, 1951); it belongs to the Gammaproteobacteria and is so far the best studied of all acidophilic prokaryotes. However, details of the flow of electrons from Fe(II) to molecular oxygen are still debated. Apparently, Fe(II) is oxidized on the cell surface at the outer membrane. Subsequently, a variety of membrane-bound and periplasmic cytochromes plus the periplasmic copper protein rusticyanin transfer the electrons to a cytochrome c oxidase. There the electrons are used together with protons to reduce molecular oxygen to water (Johnson, 2007; Castelle et al., 2008).

Neutrophilic aerobic Fe(II)-oxidizing bacteria So far, only few representatives of the Alpha-, Beta-, and Gammaproteobacteria are described to grow by dissimilatory aerobic oxidation of Fe(II) at neutral pH. This physiological group gains more energy than their acidophilic counterparts because the difference between the redox potentials of the relevant redox couples is greater at neutral than at acidic pH values (Fe(III)/Fe(II) < +100 mV; O2/H2O +820 mV; Kappler and Straub, 2005; Appelo and Postma, 2007). However, in oxic environments of neutral pH Fe(II) has a half-life in the order of few minutes and is chemically oxidized to ferric iron (Canfield et al., 2005). This competition with the chemical oxidation is likely the major reason why neutrophilic aerobic Fe(II)oxidizing bacteria thrive preferentially in microoxic niches or oxic/anoxic boundaries. Laboratory studies showed that the activity of Fe(II)-oxidizing bacteria contributes up to 90% to the formation of Fe(III) under such conditions. Typical habitats for neutrophilic aerobic Fe(II)-oxidizing bacteria include groundwater springs, freshwater, and marine hydrothermal vents, wetlands, and plant rhizospheres (Emerson, 2000). Due to their production of conspicuous iron-encrusted twisted stalks or sheaths, Gallionella ferruginea and Leptothrix ochracea were the first Fe(II)-oxidizing bacteria that were already discovered in the nineteenth century. Only during the past few years, additional morphologically inconspicuous aerobic Fe(II)-oxidizing bacteria were isolated, e.g., Ferritrophicum radicicola and Sideroxydans paludicola (Canfield et al., 2005, Weiss et al., 2007). According to physiological studies, several of the novel strains are obligate aerobic Fe(II)-oxidizing bacteria, i.e., Fe(II) is the only electron donor and molecular oxygen the only electron acceptor utilized (Weiss et al., 2007). Neutrophilic anaerobic Fe(II)-oxidizing nitrate-reducing prokaryotes Under anoxic conditions, bacteria (Alpha- Beta-, Gammaand Deltaproteobacteria) and archaea (Archaeoglobales) are capable of oxidizing Fe(II) with nitrate as the terminal electron acceptor. At pH 7, all redox pairs of the nitrate reduction pathway can accept electrons from Fe(II), because their redox potentials are more positive than that

FE(II)-OXIDIZING PROKARYOTES

of the iron redox couple (Straub et al., 1996). Nitratedependent Fe(II) oxidation was reported for diverse habitats such as rice paddy soil, activated sewage sludge, freshwater or marine sediments (Canfield et al., 2005). The first observation of this metabolism was made with a freshwater enrichment culture: Fe(II) was oxidized to Fe(III) and nitrate was concomitantly and stoichiometrically reduced to molecular nitrogen (Straub et al., 1996). As Fe(II) was the sole electron donor supplied, Fe(II) oxidation coupled to nitrate reduction definitely supported cell growth in this enrichment culture. The situation is not so clear in most pure cultures of Fe(II)-oxidizing nitrate-reducing bacteria (e.g., Acidovorax sp. strain BoFeN1 or Thermomonas sp. strain BrG3) because they need an organic co-substrate for growth. In such strains it is questioned whether Fe(II) oxidation is beneficial and supports cell growth or whether Fe(II) is just oxidized in a rather unspecific side reaction. All known species are able to utilize alternative electron donors (e.g., hydrogen, organic acids) and alternative electron acceptors (nitrite, oxygen, ferric iron). Geobacter metallireducens, isolated as Fe(III)-reducing bacterium, couples the oxidation of Fe(II) to the reduction of nitrate to ammonium; it is unclear if G. metallireducens obtains energy by this process (Lovley et al., 2004; Canfield et al., 2005). Ferroglobus placidus is so far the only representative of the domain Archaea that grows by coupling the dissimilatory oxidation of Fe(II) to the reduction of nitrate to nitric oxide and nitrogen dioxide. Alternatively, F. placidus utilizes hydrogen, sulfide, acetate, or monoaromatic compounds as electron donor and thiosulfate or Fe(III) as electron acceptor (Hafenbradl et al., 1996; Lovley et al., 2004).

Fe(II) as electron donor for assimilative reduction reactions Various groups of prokaryotes are able to reduce oxidized inorganic compounds for the synthesis of biomass. Such assimilative reduction reactions require a source of electrons (Canfield et al., 2005). Phylogenetically and physiologically different groups of chemotrophic and phototrophic prokaryotes have the ability to exploit Fe(II) as source of electrons for assimilative reduction reactions such as the fixation of carbon dioxide. Chemotrophic acidophilic and neutrophilic, aerobic and anaerobic Fe(II)oxidizing prokaryotes have been discussed before because they oxidize the major fraction of Fe(II) during respiratory energy generation. In contrast, phototrophic Fe(II)oxidizing prokaryotes utilize light as energy source and oxidize Fe(II) exclusively for assimilative purposes. The assimilative oxidation of Fe(II) leads to the formation of Fe(III) which is barely soluble at neutral pH. Neutrophilic phototrophic Fe(II)-oxidizing bacteria therefore have to cope with a virtually insoluble metabolic end product. Anaerobic phototrophic Fe(II) oxidation Anoxygenic phototrophic bacteria were the first prokaryotes discovered that oxidize Fe(II) in the absence of

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molecular oxygen (Widdel et al., 1993). So far, only few cultures of Fe(II)-oxidizing anoxygenic phototrophic bacteria were established either from freshwater or from marine sediments. They are affiliated with the purple sulfur (Thiodictyon sp. strain F4), the purple non-sulfur (e.g., Rhodobacter sp. strain SW2, Rhodovulum robiginosum), or the green sulfur bacteria (Chlorobium ferrooxidans). All strains have the ability to utilize alternative electron donors such as hydrogen, reduced sulfur compounds, or organic acids (Canfield et al., 2005). The discovery of anoxygenic phototrophic bacteria that directly catalyze the oxidation of Fe(II), provides an alternative or additional explanation for the origin of Precambrian banded iron formations (Widdel et al., 1993; Kappler and Straub, 2005).

Summary Phylogenetically diverse species of the prokaryotic domains Bacteria and Archaea have the ability to oxidize Fe(II) during aerobic or anaerobic respiration and/or for assimilative reduction reactions. Only one electron can be obtained from the oxidation of Fe(II) to Fe(III) and Fe(II)oxidizing prokaryotes have to oxidize large quantities of Fe(II) to sustain cell metabolism. Due to the complex geochemistry of iron, acidophilic and neutrophilic Fe(II)oxidizing prokaryotes have to cope with different constraints: (1) Acidophilic Fe(II)-oxidizing prokaryotes gain less energy in respiration than their neutrophilic counterparts; (2) Neutrophilic Fe(II)-oxidizing prokaryotes have to cope with a virtually insoluble metabolic end product. It is discussed that anoxygenic phototrophic Fe(II)oxidizing bacteria contributed to the generation of Precambrian banded iron formations. Bibliography Appelo, C. A. J., and Postma, D., 2007. Geochemistry, Groundwater and Pollution. Leiden: Balkema. Canfield, D. E., Thamdrup, B., and Kirstensen, E., 2005. Aquatic Geomicrobiology. San Diego: Elsevier. Castelle, C., Guiral, M., Malarte, G., Ledgham, F., Leroy, G., Brugna, M., and Giudici-Orticoni, M. T., 2008. A new iron-oxidizing/O2-reducing supercomplex spanning both inner and outer membranes, isolated from the extreme acidophile Acidithiobacillus ferrooxidans. The Journal of Biological Chemistry, 283, 25803–25811. Cornell, R. M., and Schwertmann, U., 2003. The Iron Oxides Structure, Properties, Reactions, Occurrence and Uses. Weinheim: Wiley-VCH. Emerson, D., 2000. Microbial oxidation of Fe(II) and Mn(II) at circumneutral pH. In Lovely, D. R. (ed.), Environmental Microbe–Mineral Interactions. Washington: ASM, pp. 31–52. Hafenbradl, D., Keller, M., Dirmeier, R., Rachel, R., Roánagel, P., Burggraf, S., Huber, H., and Stetter, K. O., 1996. Ferroglobus placidus gen. nov., sp. nov., a novel hyperthermophilic archaeum that oxidizes Fe2+ at neutral pH under anoxic conditions. Archives of Microbiology, 166, 308–314. Johnson, D. B., 2007. Physiology and ecology of acidophilic microorganisms. In Gerday, C., and Glansdorff, N. (eds.), Physiology and Biochemistry of Extremophiles. Washington: ASM, pp. 257–270.

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Johnson, D. B., Ghauri, M. A., and McGinness, S., 1993. Biogeochemical cycling of iron and sulphur in leaching environments. FEMS Microbiology Reviews, 11, 63–70. Kappler, A., and Straub, K. L., 2005. Geomicrobiological cycling of iron. Reviews in Mineralogy and Geochemistry, 59, 85–108. Lovley, D. R., Holmes, D. E., and Nevin, K. P., 2004. Dissimilatory Fe(III) and Mn(IV) reduction. Advances in Microbial Physiology, 49, 219–286. Straub, K. L., Benz, M., Schink, B., and Widdel, F., 1996. Anaerobic, nitrate-dependent microbial oxidation of ferrous iron. Applied and Environmental Microbiology, 62, 1458–1460. Temple, K. L., and Colmer, A. R., 1951. The autotrophic oxidation of iron by a new bacterium: Thiobacillus ferrooxidans. Journal of Bacteriology, 62, 605–611. Weiss, J. V., Rentz, J. A., Plaia, T., Neubauer, S. C., Merrill-Floyd, M., Lilburn, T., Bradburne, C., Megonial, J. P., and Emerson, D., 2007. Characterization of neutrophilic Fe(II)-oxidizing bacteria isolated from the rhizosphere of wetland plants and description of Ferritrophicum radicicola gen. nov. sp. nov., and Sideroxydans paludicola sp. nov. Geomicrobiology Journal, 24, 559–570. Widdel, F., Schnell, S., Heising, S., Ehrenreich, A., Assmus, B., and Schink, B., 1993. Ferrous iron oxidation by anoxygenic phototrophic bacteria. Nature, 362, 834–836.

Cross-references Aerobic Metabolism Anaerobic Transformation Processes, Microbiology Archaea Bacteria Banded Iron Formations Chemolithotrophy Fe(III)-Reducing Prokaryotes Gallionella Isotope Fractionation (Metal) Isotopes and Geobiology Leptothrix Metals, Acquisition by Marine Bacteria Nitrogen Photosynthesis Siderophores

FE(III)-REDUCING PROKARYOTES Kristina L. Straub University of Vienna, Vienna, Austria

Synonyms Fe-reducers; Fe-reducing prokaryotes/microorganisms; Ferric iron-reducing prokaryotes/microorganisms; Ironreducers; Iron-reducing prokaryotes/microorganisms; Iron(III)-reducing prokaryotes/microorganisms Definition Fe(III)-reducing prokaryotes. Various species of the prokaryotic domains Bacteria and Archaea have the ability to reduce Fe(III), ferric iron, to Fe(II), ferrous iron, by the transfer of electrons. The electrons for Fe(III) reduction derive mainly from the metabolic oxidation of organic compounds or hydrogen.

Introduction Reduction of Fe(III) by prokaryotes has been known since the beginning of the twentieth century but was not considered to be of importance. At that time, only few bacterial strains were known to reduce small amounts of Fe(III) during fermentation. In addition, it was misleadingly presumed that prokaryotes cause reduction of Fe(III) mainly indirectly by producing sulfide, releasing organic compounds, lowering the redox potential, or decreasing the pH. This perspective changed notedly with the discovery of dissimilatory Fe(III)-reducing bacteria, i.e., bacteria that utilize Fe(III) as terminal electron acceptor in an anaerobic type of respiration (Balashova and Zavarzin, 1979, 1980; Lovley and Phillips, 1988; Myers and Nealson, 1988). In order to gain energy for maintenance and growth by this type of respiration, significant amounts of ferric iron have to be reduced. Additionally, it became clear that neither lowering the redox potential nor decreasing the pH is sufficient to cause Fe(III) reduction. Furthermore, it was demonstrated that only very few organic compounds (e.g., fructose, cysteine) were able to reduce Fe(III) chemically at significant rates. In the last 20 years, numerous dissimilatory Fe(III)-reducing prokaryotes were identified and it is now generally accepted that this physiological group of prokaryotes has a strong influence on the iron geochemistry of most environments. Because many Fe(III)-reducing prokaryotes also have the ability to catalyze the reduction of Mn(IV) or Mn(III) to Mn(II) they additionally influence the geochemistry of manganese (Lovley et al., 2004; Canfield et al., 2005; Kappler and Straub, 2005). Evolutionary consideration The ability to reduce Fe(III) occurs widely within the domains Bacteria and Archaea. Most known Fe(III)reducing prokaryotes that grow by Fe(III) reduction belong to the phylum Proteobacteria with representatives in each of the five subdivisions Alpha- (e.g., Acidiphilium rubrum), Beta- (e.g., Ferribacterium limneticum), Gamma- (e.g., Shewanella oneidensis), Delta- (e.g., Geobacter sulfurreducens), and Epsilonproteobacteria (e.g., Sulfurospirillum barnesii). Several Gram-positive bacteria that belong to the phylum Firmicutes (e.g., Thermoanaerobacter siderophilus) and some separate bacterial lineages like Geothrix fermentans or Geovibrio ferrireducens also have the ability to reduce ferric iron (Lovley et al., 2004). Fe(III) reduction in the domain Archaea was discovered by testing pure cultures that had originally not been isolated with Fe(III): Representatives of the phyla Euryarchaeota (e.g., Pyrococcus furiosus) and Crenarchaeota (e.g., Pyrobaculum islandicum) showed reduction of ferric iron (Vargas et al., 1998). The great phylogenetic diversity of Fe(III)-reducing prokaryotes with representatives in all deeply branching phyla, along with iron isotope studies, supports the hypothesis that Fe(III) reduction was an early mode of energy metabolism. Further geochemical evidences

FE(III)-REDUCING PROKARYOTES

suggest that other major electron acceptors such as sulfate, nitrate, or molecular oxygen were not available on early Earth (Lovley et al., 2004; Canfield et al., 2005; Johnson et al., 2008).

Geochemical aspects of Fe(III) Iron is the fourth most abundant element in the Earth’s crust and Fe(III) minerals may account for a few percent dry weight of rocks, soils, and sediments. Hence, in most anoxic ecosystems Fe(III) minerals are the dominant electron acceptors for bacteria and archaea. Today, 16 different Fe(III) oxides, hydroxides, or oxide hydroxides are known and for simplicity they are often collectively termed Fe(III) oxides. They are all composed of Fe together with O and/or OH and differ in composition and crystal structure. Widespread Fe(III) oxides include ferrihydrite, goethite, hematite, lepidocrocite, and magnetite (Cornell and Schwertmann, 2003). In contrast to other electron acceptors such as oxygen, nitrate, or sulfate, Fe(III) oxides are poorly soluble at neutral pH. The solubility of Fe(III) oxides at neutral pH depends furthermore on the type of oxide, the degree of crystallinity and the crystal size, and may range from 10
9 M (e.g., ferrihydrite, low crystallinity, small crystals) to 10
18 M (e.g., goethite, high crystallinity, large crystals). At circumneutral pH, Fe(III)-reducing prokaryotes therefore have to cope with a virtually insoluble electron acceptor. With increasing acidity, the solubility of Fe(III) oxides increases and ferric iron is well soluble at pH values below 2.5 (Cornell and Schwertmann, 2003). Concomitant with the change in pH and Fe(III) oxide solubility, the redox potential of the transition between ferric and ferrous iron changes significantly. The redox potential of the redox pair Fe3þ/Fe2þ is þ770 mV only at acidic pH values. At neutral pH, with poorly crystalline Fe(III) oxides (ferrihydrite, lepidocrocite) as the oxidant, the redox potential ranges between þ100 and
100 mV, while the redox potential of more crystalline Fe(III) oxides (goethite, hematite, magnetite) may be as low as
300 mV (Canfield et al., 2005; Straub et al., 2001). In respect of solubility and energetics, acidophilic and neutrophilic Fe(III)-reducing prokaryotes cope with entirely different substrates. Reduction of Fe(III) by fermenting bacteria The first Fe(III)-reducing prokaryotes that were studied in laboratory cultures belonged to the domain Bacteria and reduced only small amounts of Fe(III) during fermentative growth on organic substrates such as glucose or malate. This physiological group comprises of a variety of bacteria and includes anaerobes (e.g., Clostridium pasteurianum, Vibrio spp.), facultative anaerobes (e.g., Escherichia coli, Paenibacillus polymyxa), and aerotolerants (e.g., Lactobacillus lactis, Lactococcus lactis). Accordingly, Fe(III) reduction by fermenting bacteria was observed under both anoxic and oxic culture conditions. For fermenting bacteria, Fe(III) reduction is

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only a minor pathway for electron disposal and research on this type of metabolism fell behind efforts to understand dissimilatory Fe(III) reduction. The mechanism(s) of electron transfer to Fe(III) during fermentation is unknown and it is also unclear whether this process yields energy for growth. Thermodynamic calculations suggest that Fe(III) reduction during fermentation results in a slightly greater energy yield than fermentation alone but experimental evidences are ambiguous (Lovley, 1991; Canfield et al., 2005). So far no representative of the domain Archaea was described to reduce Fe(III) during fermentative growth.

Dissimilatory Fe(III)-reducing prokaryotes Dissimilatory Fe(III)-reducing prokaryotes utilize Fe(III) as terminal electron acceptor in anaerobic respiration, i.e., they gain energy by coupling the oxidation of an electron donor to the reduction of Fe(III). As in the reduction of Fe(III) to Fe(II) only one electron can be disposed, dissimilatory Fe(III)-reducing prokaryotes have to reduce large quantities of Fe(III) to gain enough energy for maintenance and growth. Numerous strains of phylogenetically dispersed, dissimilatory Fe(III)-reducing prokaryotes have been isolated into pure culture or have been detected by molecular techniques in a wide range of pristine or contaminated habitats, including freshwater and marine sediments, wetlands, soils, aquifers, the deep subsurface, deep subterranean thermal water, hydrothermal vents, and hot springs (Lovley et al., 2004). This widespread occurrence correlates well with the ubiquitous presence of Fe(III) minerals. Consistent with the great variety of ecosystems inhabited, pure cultures of Fe(III)-reducing prokaryotes show a wealth of physiological qualities in respect to oxygen concentration (strict or facultative anaerobes), temperature (growth reported between 4 and 121 C), pH (growth reported between pH 1 and 9), and salinity (freshwater and marine species). In addition, autotrophic growth with carbon dioxide as sole carbon source and fixation of molecular nitrogen was described for some species of Fe(III)-reducing prokaryotes. No prokaryote characterized so far, depends solely on Fe(III) as terminal electron acceptor for growth. In the absence of Fe(III), prokaryotes either grow by fermentation (e.g., Thermotoga maritima) or utilize alternative electron acceptors. Common alternative electron acceptors include oxygen (e.g., Pantoea agglomerans), nitrate (e.g., Geobacter metallireducens), manganese (e.g., Deferribacter thermophilus), reduced sulfur compounds (e.g., Desulfuromusa kysingii), and fumarate (e.g., Desulfuromonas acetexigens). Furthermore, some species transfer electrons to heavy metals (e.g., uranium, chromium), graphite electrodes, humic substances, or chlorinated compounds (e.g., tetrachloroethene, trichloroacetic acid). The vast majority of Fe(III)-reducing prokaryotes utilizes fermentation end products such as acetate, ethanol, or hydrogen as electrons donors. Only few prokaryotes are known to couple the dissimilatory reduction of Fe(III) to the oxidation of sugars (e.g., Acidiphilium

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FE(III)-REDUCING PROKARYOTES

cryptum), amino acids (e.g., Geothermobacter ehrlichii), long-chain fatty acids (e.g., Desulfuromonas palmitatis), monoaromatic compounds (e.g., Geobacter metallireducens), or reduced inorganic sulfur compounds (e.g., Acidithiobacillus ferrooxidans).

Acidophilic Fe(III)-reducing bacteria A variety of autotrophic and heterotrophic bacteria that thrive in environments of low pH (6 h

Fixation of cellsb
20°/+4°Cb 0.5 h to 96 h

Yes/partly No No No No

Partly Yes Yes Yes Yes

Only with multiplex PCR No (for DNA) No No Yes Yes Yes (for qPCR) Possible Inhibition >5 gene copies

Yesc Partlyd Yes Yes Partly Noe No Possible Autofluorescence 105–7 cells/mlf

a

Including different FISH protocols and advanced microscopy tools. In certain cases, samples may be stored at
80°C and the fixation omitted (Yilmaz et al., 2010b). c Only with certain FISH protocols like mRNA-CARD FISH, RING-FISH, gene-FISH, and in situ RCA. d Depends on the FISH protocol and how this is combined with other analytical tools and appropriate references. e Exception, FISH protocols based on PCR. f Depends on concentration/diluation procedure. b

of the samples as well as the morphological integrity of cells during the different hybridization steps (elevated temperature, exposure to enzymes, detergents and osmotic gradients) and prevent cell losses, while simultaneously making the cells permeable for the probe(s) (Amann, 1995).Unfortunately, there is not one plain fixation protocol for all cell types for all different FISH protocols. Different fixation protocols have been developed to cope with the complexity of archaea, bacteria, and Eukarya with regard to their cell envelope packages and the type of probes to be employed for a particular FISH protocol. A suboptimal fixation protocol may make cells impermeable for probes or even cause cell lysis, especially during longer storage times or when exposing the cells to certain procedures such as incubation with enzymes prior to the FISH procedure. Apart from the selection of an optimal fixative reagent, it is also relevant to pay attention to the concentration of the fixative reagent and the incubation time and conditions. Fixation times may range from a few min up to >24 h, depending on cell type and FISH protocol, and are generally followed by a washing step for removal of the fixative reagent. The two most common fixation procedures are based on either aldehydes (e.g., formalin, paraformaldehyde (PFA), glutaraldehyde) or on alcohols (e.g., ethanol, methanol). In the

majority of cases, Gram-negative cells within the domain Bacteria are usually fixed with 4% PFA, and Gram-positive cells within the domain Bacteria with ice cold 96% ethanol (Amann, 1995). To a certain extent, some Gram-negative species will also tolerate EtOH fixation, and some Gram-positive species (general exceptions are, e.g., Actinomycetes and Streptococci) may also tolerate PFA fixation (especially if the incubation time is short, see, e.g., de los Reyes et al., 1997). The previously mentioned fixation protocols may also work well for certain Archaea and Eukarya species, but in some cases, especially for certain Archaea species (Nakamura et al., 2006) or eukaryotes like fungi (Baschien et al., 2001; Tsuchiya and Taga, 2001; Weber et al., 2007) or protozoa, see, e.g., Fried et al. (2002); Petroni et al. (2003); Fried and Foissner (2007); Weber et al. (2007), other fixative reagents or post treatments with enzymes like chitinase or endo-isopeptidase are more suitable. When dealing with unknown samples, it is therefore recommended to explore the suitability of different fixation protocols. Irrespective of the fixation protocol used, the cells must be fixed as soon as possible with optimally prepared fixation solutions, preferably already during the sampling occasion to avoid population shifts and for optimal evaluation of true probe signal intensities as these

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FLUORESCENCE IN SITU HYBRIDIZATION (FISH)

may in certain cases also serve as an indicator of activity (see, e.g., Wagner et al., 1995; Molin and Givskov, 1999; Rossetti et al., 2007). 3. Storage of fixed cells: Fixed samples (in wet or dry state, such as on a microscope slide or on a membrane filter) can be stored at room temperature or at +4°C for shorter periods. For optimal long-term storage at
20°C, fixed cells are generally dissolved in a 1:1 (v/v) solution with a 50% end concentration of ethanol in order to avoid freezing (Amann et al., 1994). Cells concentrated on a membrane filter are normally stored dry at
20°C (e.g., Teira et al., 2004). So far, no systematic studies on the maintenance of cell integrity or ribosome content in fixed cells after longer storage periods exist, but it is generally recommended that once the cells have been fixed, quantitative studies should be performed within days to avoid biases caused by unknown deterioration of cells or decrease of ribosome content.

Preparations for omics-based applications For genomic or proteomic applications, unfixed biomass is desirable to minimize possible sequencing biases introduced by cross-linking and to avoid possible cell lysis difficulties, so that FISH targeted cells can be enriched by, e.g., flow cytometry and subjected to, e.g., whole genome application. Recently, Yilmaz et al. (2010b) demonstrated that the standard oligonucleotide FISH protocol can also be performed on at least two different types of frozen, non-fixed complex environmental samples using Archaea as well as Bacteria targeted probes. However, more research is needed to explore further possibilities as well as limits of this approach, such as if this is quantitative and if this applies to any kind of cells types or other environmental samples, irrespective of growth and storage conditions, as well as if this approach is also suitable for other types of FISH protocols. FISH for in situ detection of ribosomal genes General background: The ribosomal gene was and will at least for the near future remain the standard gene for reconstruction of phylogeny, despite the growing amount of genome sequences which is presently revolutionizing our current concepts of taxonomy, phylogeny, and development of comprehensive biomarkers for improved identification (see, e.g., Konstantinidis et al., 2009). The main benefits of the ribosomal gene is that it is a functionally conserved molecule present in high numbers in all organisms from all domains and is composed of sequence regions of higher and lower evolutionary conservation. The variable conservation degree makes it particularly ideal for design of biomarkers for nucleic acid–based detection and quantification of different levels of microbial diversity (Ludwig et al., 1998; Pruesse et al., 2007; Amaral-Zettler et al., 2008; Yarza et al., 2008). Although several different types of ribosomal subunits exist, only a few of these are currently used as targets for probes in

FISH studies. For most FISH studies, the main gene target is the 16S rRNA gene for prokaryotes, and the 18S-rRNA gene for eukaryotes (Ludwig et al., 1998; Metfies et al., 2008). However, other multicopy ribosomal genes or even stable multicopy RNA molecules have been (5S-rRNA, tmRNA), or may be occasionally used as alternative targets for FISH probes (23S-rRNA, 28S-rRNA, precursor rRNA, intergenic spacer region. These alternative gene targets may overcome some of the problems occasionally encountered when targeting the 16S rRNA gene, such as low information content and limited probe accessibility or information about activity status (Trebesius et al., 1994; Oerther et al., 2000; Schmid et al., 2001; Schonhuber et al., 2001; Zimmermann et al., 2001; Peplies et al., 2004). However, despite this, the 16S rRNA gene has remained the main gene target in FISH applications, especially for environmental systems with unknown diversity, since the 16S rRNA gene is nowadays routinely sequenced in most studies so that the amount of retrieved 16S rRNA gene sequences has thus come to outrange all other databases of other gene sequences. Principle of FISH: The FISH procedure generally consists of the following steps (Adapted from Manz et al., 1992 and Amann, 1995): 1. Optional: For certain cases an additional posttreatment of the fixed/non-fixed biomass may be needed for increased permeabilization of probes through rigid cell envelope packages, or for removal of confounding background, by, e.g.:  Incubation with enzymes like lysozyme, protease, proteinase K, mutanolysine, endoisopeptidase, chitinase  Removal of wax or chemical precipitates by chemical solvents  Use of detergents (SDS) or other chemicals like hydrochloric acid  Physical treatment like exposure to microwave radiation  Addition of chemicals to avoid dissolution of essential compounds 2. Successive dehydration in an EtOH series (50%, 80%, 96%, each step between 1–3 min). 3. Hybridization of the fixed sample with the probe on a microscope slide or in solution under stringent conditions (Figure 1). Stringent conditions are achieved by keeping the hybridization temperature constant (+46°C) but varying the formamide concentration for different probes (Stahl and Amann, 2001). 4. Subsequent washing with preheated buffered washing solutions at a slightly higher temperature (+48°). After this, the hybridized samples can be evaluated or stored at dark prior to evaluation (for a shorter period at room temperature or at +4°, for longer periods, at
20°C). 5. Optional: Counter stain with, e.g., an appropriate nucleic acid stain like DAPI (4′,6-diamidino-2-phenylindole) or SYBR Green (e.g., Wagner et al., 1994, or Engel et al., 2003).

FLUORESCENCE IN SITU HYBRIDIZATION (FISH)

Fixed/nonfixed sample

Ethanol series dehydration (50%, 80%, 100%)

Drying

377

Hybridization buffer with fluorescently-labeled phylogenetic probe(s)

Hybridization

Drying

Washing step, 15 min, 48⬚C, water bath Incubation at 46⬚C in a humid chamber Stringent

Embedding with antifading reagents, coverslip, fluorescence Embedding microscopy

washing Pulp soaked with hybridization buffer

a

Ribosomes

Nucleoid

Cytoplasma

Nucleoid FISH

b

Cell wall cytoplasmic membrane

Ribosomes targeted by fluorescent probes

Fluorescence In Situ Hybridization (FISH), Figure 1 (a) Schematic outline of the standard oligonucleotide FISH procedure on microscope slides. (b) Schematic outline of a cell before and after the standard oligonucleotide FISH procedure.

6. Evaluation is achieved by: Visualization:  Standard epifluorescence microscopy, or confocal laser

scanning microscope for three-dimensional reconstruction (Wagner et al., 1998).  Combination with other microscopes like electron microscopy (e.g., Gerard et al., 2005), RAMAN spectroscopy combined with a microscope (Wagner, 2009), or nanoSIMS (Behrens et al., 2008; Orphan, 2009). For digital image analysis of FISH images, various software packages are available, such as CMEIAS, COMSTAT, and daime (Heydorn et al., 2000; Daims and Wagner, 2007; Zhou et al., 2007; Daims, 2009). Other options:  Membrane-based evaluation (e.g., dot blot, Wagner

et al., 1994).  Combination with other analytical tools (e.g., screening

of uptake of radioactive tracers or stable isotopes

(Nielsen et al., 2005; Orphan, 2009) or other FISH protocols targeting non-ribosomal genes or their transcripts for exploring activities or functions.  Reverse hybridization by microarray technology, where multiple probes are deposited on a support material to hybridize a labeled sample with the probes (DeSantis et al., 2005; Loy and Bodrossy, 2006; Taylor et al., 2010).  Cell sorting or probe signal intensity evaluation by flow cytometry (Behrens, 2003; Amann and Fuchs, 2008), RING (Recognition of IGdividual Genes)-FISH (Zwirglmaier et al., 2004b), or magneto-FISH (Orphan, 2009).  Whole genome amplification (Orphan, 2009; Yilmaz et al., 2010b). Basic equipment and reagents for standard FISH applications:  Hybridization oven (46°C)  Water bath (48°C)  Polypropylene screw top tubes, filter paper

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 Reagents (for chemical composition, check, e.g.,

  





Amann, 1995): – NaCl – Tris/Phosphate buffer – EDTA – SDS – Formamide – 50%, 80%, 96% EtOH – Reagents/equipment for improved probe accessibility Labeled probes Counterstains like nucleic acid stains (e.g., DAPI (4′,6diamidino-2-phenylindole) or SYBR Green) Microscope slides (possibly coated with, e.g., polyL-lysine (Sigma-Aldrich, www.sigmaaldrich.com/sigmaaldrich/home.html) or a gelatine-chromium sulphate solution (Amann, 1995) Mounting medium of appropriate pH (e.g., from Citifluor, www.citifluor.co.uk/; Vectashield, www. vectorlabs.com/; various mounting media from Invitrogen, www.invitrogen.com/site/us/en/home.html) Some form of fluorescence microscope.

FISH probes (probe design, probe evaluation, probe databases) For a proper interpretation of FISH experiments, probes need to be optimally designed and handled. Several different types of softwares (e.g., the ARB software package, (www.arb-home.de/) and Primrose (www. bioinformatics-toolkit.org/index.html)) and detailed guidelines for probe design for particularly FISH experiments exist (Amann, 1995; Amann and Schleifer, 2001; Stahl and Amann, 2001), and (Hugenholtz et al., 2001). Below follows an updated summary of some of those guidelines, focusing mainly on applications with the ARB software package: 1. Construction of a molecular data base: Prior to all probe design, a comprehensive data base of all relevant complete gene sequences of the target group and relevant nontarget groups should be collected for a thorough evaluation of phylogenetic relationships. Partial sequences may also be included, but these should be handled more critically due to their limited value for a full evaluation of all potential probe targets. Sequences can be downloaded from different categories of databases: (1) all kinds of gene sequences: EMBL (www.embl.de/), NCBI (www.ncbi.nlm.nih. gov/), (2) regularly updated and curated ribosomal gene sequences: RDP (http://rdp.cme.msu.edu/), SILVA (www.arb-silva.de/), Green Genes (http:// greengenes.lbl.gov/cgi-bin/nph-index.cgi); the ribosomal internal spacer collection (http://egg.umh.es/ rissc/); (3) functional genes: Fun Gene (http:// fungene.cme.msu.edu/index.spr). Principally, most softwares that can align sequences, perform phylogenetic evaluations, and design biomarkers like PCR primers can be used for FISH probe design as well.

However, the bioinformatic software package ARB (www.arb-home.de/) using the curated SILVA database (www.arb-silva.de/) is highly recommended, as several useful and partly unique tools have been implemented into this package to facilitate a straightforward and advanced design, evaluation and visualization of probes and their target (Ludwig et al., 2004; Kumar et al., 2005; Kumar et al., 2006; Pruesse et al., 2007). 2. Hierarchic probe design strategy: Where possible, it is recommended to design a hierarchic probe set consisting of several probes targeting different phylogenetic levels of the target organism/group (see, e.g., Amann and Schleifer, 2001; Amann and Fuchs, 2008). If the different probes are labeled with different fluorochromes and combined in one FISH experiment on a mixture of cells with different phylogenetic relationships, different overlaps of the probes will produce different color combinations depending on the taxonomical relationships. This facilitates a straightforward top to bottom identification of the taxonomical status (e.g., from phylum down to species level) of different cell species in a complex sample (Amann, 1996). 3. General rules for standard oligonucleotide probe design: (a) Select the appropriate gene sequences of the desired target group. Depending on bioinformatic software, design the probe according to the recommended procedure. For example, if using the ARB software (Ludwig et al., 2004), select from the menu “probe design” an appropriate PT server, then define appropriate parameters such as length of probe (average length of probes for standard oligonucleotide probes is between 15–25 nucleotides), minimum group and maximum nongroup hits, GC content, melting temperature range, range of nucleotide positions, and maximum hairpin bonds. (b) The selected probe should be completely specific (complementary) to a region of the selected target sequences and have at least one mismatch to the same target region in nontarget sequences. Optimally, the mismatch should be centralized in the nontarget sequences in order to maximize the destabilizing effect of the mismatch. (c) Perform probe match to evaluate the specificity of the probe (e.g., in the bioinformatic software package ARB (www.arb-home.de), the RDP Probe Match tool (http://rdp.cme.msu.edu/probematch/ search.jsp), the probeBase probeCheck tool (http://131.130.66.200/cgi-bin/probecheck/content. pl?id=home) or general data bases like the National Center for Biotechnology Information (www.ncbi. nlm.nih.gov/). (d) Unfortunately, there is no such thing as a perfect probe that only targets the target group as the course of the evolution of gene sequences is a random process. During a recent review Amann and Fuchs (2008) showed that it is more advisable to

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evaluate group coverages as the percentage of group members that can be identified relative to the total number of members in the target group. With this approach, three groups can be distinguished: true positives; false negatives; and false positives. Parallel to the constantly growing amount of gene sequences, the coverage of each probe target group is ultimately going to change and possibly demand more complex probe sets. For example, in the early days of the FISH technology, only one probe was needed to target all recognized species in the Bacteria domain (Amann et al., 1990). However, a decade later, Daims et al. (1999) showed that a mixture of three different Bacteria domain targeting probes were needed to cover all so far (up till 1999) recognized sublineages. (e) Evaluate the melting temperature of the probes (e.g., as described by (Hugenholtz et al., 2001) or the probeBase, www.microbial-ecology.net/ probebase/). Optimally, the probe should have a melting temperature > 57°C. If the melting temperature is < 57°C, the temperature can be increased by shifting the position of the probe or expanding the length of the probe. (f) Check probe accessibility. A reoccurring problem is the probe accessibility. The general explanation for is that certain regions of the secondary structure of the ribosomal gene are inaccessible for probes. Unfortunately, the accessible and inaccessible regions of the ribosomal molecules appear to vary from species to species, so that only precise estimations of probe accessibilities can be made for those species where a systematic probe accessibility mapping has been performed (see, e.g., Fuchs et al., 1998; Behrens, 2003a, 2006b). The general recommendation is therefore to design probes on different target positions and then to simply explore on a trial-and-error experimental basis which of the selected probes produce high probe signals. However, Yilmaz et al. (2006) showed that principally all positions on the ribosome molecule may yield satisfying probe signals if a sophisticated approach based on a thermodynamics-based probe design is employed. With this approach, FISH experiments of any given probe can be mathematically simulated. Based on this, theoretical predictions on, e.g., the probe accessibility, the formamide denaturation profile, and the potential of mismatch discriminations can be made (for further details see Yilmaz et al., 2010a). Based on this, further optimizations of the probe accessibility can be made such as modifying the length of the probe or the experimental hybridization conditions (e.g., increase of the hybridization time up to 96 h). (g) Check for potential self-complementarity of the probe which may initiate duplex formations between the probe and the target (e.g., caused by hairpins or dimers).

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(h) Naming of probe. There are three types of nomenclatures for probes: (1) the short name (e.g., EUB 338), which is often constructed by a shortage of the name of the target group (e.g., EUB for Eubacteria), followed by a number that denotes the first position of the probe on the target gene (e.g., nucleotide position 338 on the E. coli gene); (2) a name based on a comprehensive nomenclature to denote main features of the probe such as the target gene, target group, target group level, 3’end of the probe, and probe length (Alm et al., 1996); and (3) the probeBase accession number (e.g., pB-00159 for probe EUB 338) as assigned by the online probe database ProbeBase (www.microbialecology.net/probebase/). 4. Labeling of probes: The first rRNA-targeted oligonucleotide probes for FISH were radioactively labeled, developed by microautoradiography and evaluated by light microscopy (Giovannoni et al., 1988). Unfortunately, the work with radioactively labeled probes restricted the applicability of FISH. However, the applicability of FISH increased as soon as the first fluorescently labeled rRNA-targeted oligonucleotide probes were reported (DeLong et al., 1989). Probes are nowadays generally covalently labeled at the 5’end of the oligonucleotide, and the most commonly used fluorochromes for FISH are sulfoindocyanine fluorochromes like Cy3 and Cy5, fluorescein, ALEXA probes, rhodamine, and Texas red. The criteria for selecting appropriate fluorochromes are based on the capabilities of the available epifluorescence microscope and on the nature of the cells and their surrounding environment, where the aim is to avoid a confounding background and unspecific binding of fluorochromes to nontarget cells, and/or to minimize bleaching of probe signals. During the last decade, several different approaches have been used to increase the probe signal intensities in those cases where standard labeled probes produce too weak fluorescent signals. A simple strategy was to employ mixtures of monolabeled probes targeting different sites of the ribosomal gene (e.g., Amann et al., 1990; Krumholz et al., 1990; Morris et al., 2002; Sunamura and Maruyama, 2006). Another strategy is to employ alternative labeling procedures of oligonucleotide probes, such as polynucleotide FISH (e.g., Trebesius et al., 1994), horseradish peroxidase (employed in catalyzed reporter deposition (CARD-FISH), e.g., Pernthaler et al., 2002a; Wendeberg, 2010) or various types of multiple labeling of probes (Amann and Schleifer, 2001) such DOPE-FISH (Stoecker et al., 2010). 5. Purchase and handling of probes: Probes are chemically synthesized and can be purchased with or without label. In earlier times, probes had to be manually labeled (Amann, 1995), but nowadays labeled probes can be easily purchased from most providers (e.g., Biomer, Invitrogen, MWG, SIGMA) of molecular reagents like PCR primers at a rather low cost (starting

380

FLUORESCENCE IN SITU HYBRIDIZATION (FISH)

from approximately 50 USD). Probes are normally shipped in lyophilized stage and must be dissolved in either sterile water or an appropriate buffer (depending fluorochrome chemistry) to a stock solution of normally 100 pmol/µl. It is recommended to store aliquots of working solutions in individual tubes in dark at
20°C, and to avoid excessive freeze-thawing of the aliquots. For long time storage or transport at room temperature it is recommended to lyophilize the probes. 6. Evaluation of probes: (a) For any purchased probe, it is recommended to check for the probe’s concentration, length, homogeneity, and the completeness of the labeling, to avoid unwanted competition between labeled and unlabeled probes for target sites as this may reduce the probe signal intensity. This can be done as described by Stahl and Amann (2001), or as recommended by the provider who performed the labeling. (b) Newly designed probes are generally first tested under non-stringent conditions (i.e., by excluding formamide in the hybridization buffer). If positive probe signals are obtained in reference strains as expected, stringent conditions of the probe should be determined by an increasing formamide series, as described in Amann and Schleifer (2001). Probe signal intensities can be quantified by digital image analysis with, e.g., the daime software, as described by (Daims et al., 2009). (c) Combination of probes. The great advantage of FISH over standard PCR applications is that several differently labeled gene probes can be simultaneously combined and thus enable a simultaneous identification of different probe-targeted species within a complex sample. The general guidelines for optimal combination of different probes are:  Probes with identical formamide concentrations for optimal stringencies (at which the probe hybridizes to the specific target but not to nontarget groups) can be combined in the same hybridization experiment.  Probes with different formamide concentrations for optimal stringencies cannot be combined in the same hybridization experiment. Instead, successive hybridizations must be performed for each probe, starting with the probe with the highest formamide concentration. (d) Most standard epifluorescence microscopes are equipped with filter sets that allow separate recording of at least two to three differently labeled probes and one UV-based nucleic acid stain like DAPI (4′,6-Diamidino-2-phenylindol). Such recorded images can then be merged into one image with image analysis software packages. However, recent developments based on, e.g., advanced confocal laser scanning microscopes with multichannel image analysis, special tools such as spectral imaging or FISH protocols employing quantum dot

labeled probes allow the simultaneous combination of a considerably higher amount of differently labeled probes (e.g., Ainsworth et al., 2006; Anderson, 2010; Bentolila et al., 2006). 7. Probe bases: Different attempts to construct databases with FISH probes have been made during the last two decades. However, the probeBase (www.microbialecology.net/probebase/) is currently the most regularly updated data base for ribosomal targeted probes, encompassing 2,568 different probes targeting species within Archaea, Bacteria, and, to some extent, Eukarya (further probes targeting protists and cyanobacteria can be retrieved from www.sb-roscoff.fr/Phyto/Databases/RNA_probes_introduction.php) for FISH as well as for ~16 different microarray applications (version September 2010). Here, many of the so far published probes can be easily searched by different criteria (target organism, target sites, probe name, probe sequence, probeBase accession number, references). Detailed information about the probe chemistry are listed and links are provided for online probe matches to other databases like the ProbeCheck http://131.130.66.200/cgi-bin/probecheck/content.pl? id=home, Green Genes (http://greengenes.lbl.gov/cgibin/nph-index.cgi), or RDP (http://rdp.cme.msu.edu/). Furthermore, up to 150 ribosomal gene sequences retrieved from own clone libraries can be uploaded to screen for probes targeting these sequences (Loy et al., 2007, 2008). 8. Controls: For a valid evaluation of probe signals, three different categories of controls are needed: (1) Positive controls where probes are employed that should or should not overlap with the intended target gene. (2) Negative controls, to explore the impact of confounding background caused by, e.g., autofluorescence, and of nonspecific binding of fluorochrome or probe to target cells as well as nontarget cells. To achieve this, two different negative controls are recommended: exclusion of probes during the hybridization experiment, and application of negative reference probes like the NONSENSE probe (Loy et al., 2002) or the non-EUB probe (reverse complementary to the EUB probe, (Wallner et al., 1993). (3) All experiments should be performed on appropriate optimally fixed strains carefully selected to serve as positive as well as negative controls. 9. Other probe types: Note that some of the rules applied for design and handling of standard oligonucleotide probes may not be applicable to other probe types, like polynucleotide probes, PNA FISH probes, or CARD FISH probes.

General guidelines for evaluation and presentation of FISH results For optimal interpretation of FISH experiments, several parameters should be considered, as outlined below. Omitting any of these principles may lead to false interpretations of FISH results.

FLUORESCENCE IN SITU HYBRIDIZATION (FISH)

Checklist for optimal FISH experiments and evaluation of results: 1. Were optimal sampling strategies, fixation protocols, and storage conditions employed? 2. Were appropriate reference strains used simultaneously in parallel with the FISH experiments? If strains are lacking, clones based on CLONE-FISH (Schramm et al., 2002) may also be used to experimentally evaluate the FISH experiments. 3. Were the microscopical investigations performed with optimal microscope settings? 4. Were appropriate probes/probe combinations (positive as well as negative controls) used under stringent conditions? (a) If the probe signals were positive, explore if nonspecific probe signals can be unequivocally excluded or if the identity of the probe signal can be confirmed with other measures. (b) If the probe signals were negative on the sample subject to investigation but otherwise positive with positive controls, explore if this is caused either by cell losses during the hybridization procedure, low cell numbers in the original sample, or low activity (and thus low ribosome content and probe signal intensity). 5. If quantifications of FISH images were performed, explore if the amount of sample applied to the microscope slide was appropriate. A too high or too low cell concentration may result in an over- or underestimation of cell numbers. Evaluate also if the amount of microscope slide fields investigated was high enough to ensure statistical evaluations of error ranges. For further evaluation of quantification, spiking of sample may be recommendable (SPIKE-FISH, Daims et al., 2001). 6. When presenting FISH results, it is recommended to describe results as “cells targeted by probe X,” instead of uncritically identifying the probe signals with the intended target group of the probe. This is especially important when applying probes to samples with unknown microbial composition and the applied probe is known to target false-positive as well as falsenegative species and no further means have been undertaken to confirm the identity by, e.g., appropriate combinations of hierarchic FISH probe sets or subsequent sequencing (Amann and Fuchs, 2008).

Drawbacks of oligonucleotide FISH and the benefits of novel FISH technologies The first generation of FISH protocol based on oligonucleotide probes revolutionized nearly all disciplines in microbiology. However, along with applications of this protocol on different ecosystems and species, many of the limits of the first-generation FISH protocol became obvious: 1. Successful detection dependent on ribosome content (minimum 370 ± 16S rRNA molecules per E. coli cell (Hoshino et al., 2008).

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2. Confounding background caused by auto-fluorescing compounds like photosynthetic pigments, proteins like cofactors (e.g., F420 for methanogens), other parameters in the surrounding environment (e.g., precipitates, fibers, minerals, humic substances); or by unspecific absorption of fluorochrome label. 3. Probe technical problems (e.g., probe permeabilization problems, probe accessibility in the ribosomal gene, probe specificity). 4. Limited information content of the 16S rRNA gene for taxonomical resolution of closely related species. 5. Limited possibilities to confirm true identity and explore phylogenetic relationships and identify novel gene groups or species. 6. Limited information on in situ physiology in terms of: (a) Activity status (high ribosome content may reflect exponential growth stage, but this appears to be species dependent and needs to be explored for each particular species group (Wagner et al., 1995; Molin and Givskov, 1999; Rossetti et al., 2007); (b) Function, since the ribosomal gene does not reveal information about function and since the genotype is not always linked to the phenotype (e.g., many functional microbial groups are polyphyletic, such as methanogens, sulphate reducers, denitrifiers, cyanobacteria, spirochaetes, reductive dechlorinators); (c) Limited possibilities to monitor gene expressions (mRNA transcripts) or proteins. 7. Applications limited to prokaryotic and eukaryotic cells. However, the first notes on successful FISH applications on bacteriophages (personal communication, K. Zwirglmaier, D. Scanlan, A. Millard, University of Warwick, UK) was recently made with the Recognition of individual genes [RING]-FISH protocol (Zwirglmaier et al., 2004b). To overcome these limitations, several different strategies have been employed, ranging from modifying different parts of the procedures of the first-generation FISH protocol, searching for alternative gene targets, probe chemistries and labeling procedures, to combining FISH with other markers (e.g., activity stains, enzymes, isotopes), analytical procedures, and advanced microscopes or other analytical tools for evaluation of several different parameters for a holistic interpretation of in situ microbial ecology. Today, at least four categories of advanced FISH strategies can be recognized, which can be combined in a multitude of ways for various novel applications. Category I – increasing the probe signal intensity:  Incubate sample prior to fixation with substrates or

stress-promoting compounds like chloramphenicol (Kalmbach et al., 1997; Ouverney et al., 1997).  Employ non-fluorescent labeling of probes for increased detection of cells in, e.g., a strongly autofluorescing background based on, e.g., radiolabels (Giovannoni

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et al., 1988); or enzymes (Zarda et al., 1991; Amann et al., 1992; Lim et al., 1993). Employ alternative probe design based on helper probes for oligonucleotide probes to increase probe accessibility (Fuchs, 2000). Employ alternative probe chemistries for increased hybridization efficiencies and specificities based on peptide nucleic acid probes (PNA) (Perry-O’Keefe et al., 2001), or locked nucleic acid (LNA) probes (Kubota et al., 2006a). Explore probe accessibilities and thermodynamicsbased probe design to predict probe accessibilities and optimal hybridization conditions (Yilmaz et al., 2006; Yilmaz et al., 2010a), and/or in nucleotide or probe quenching (Behrens, 2003; Behrens et al., 2004; Wagner et al., 2003). Employ self-ligating probes to overcome background autofluorescence and avoid washing steps (Sando and Kool, 2002). Employ PCR-generated probes to generate polynucleotide probes (Trebesius et al., 1994; Zimmermann et al., 2001; Meisinger et al., 2007; Pernthaler, 2002; Preston et al., 2002; Moraru et al., 2010). Employ modified fluorescent labeling technologies based on horse radish peroxidase labeled probes and catalyzed reporter deposition (CARD)-FISH (Pernthaler et al., 2002a, b; Hoshino et al., 2008; Pavlekovic et al., 2009); multiple labeling of probes (Amann and Schleifer, 2001; Stoecker et al., 2010) or quantum dot conjugated probes (Bentolila et al., 2006).

Category II – exploring activity or function:  Combine FISH with rRNA dot slot to quantify cellular

rRNA content (Ravenschlag et al., 2000).  Employ FISH probes targeting the precursor 16S rRNA 







gene and intergenic spacer regions (Oerther et al., 2000; Schmid et al., 2001; Moter et al., 2010). Combine FISH with different activity indicating stains like redox dyes (Nielsen et al., 2003), halogenated thymidine analogue bromodeoxyuridine (BrdU) (Pernthaler et al., 2002), exo-enzyme labeled probes (Xia et al., 2007), or antibodies to target cell surface markers or proteins (Currin et al., 1990; Lubeck et al., 2000; Gmür and Lüthi-Schaller, 2007). Incubate sample with radioactively labeled substrates or stable isotopes to explore specific uptake of labeled substrates by micro-autoradiography (Nielsen et al., 2005), RAMAN spectroscopy (Wagner, 2009), or nano-SIMS (Behrens et al., 2008; Orphan, 2009). Target functional genes or even the mRNA gene by CARD-FISH (Wagner et al., 1998; Pernthaler and Amann, 2004; Orphan, 2009; Pilhofer et al., 2009), two-pass TSA-mRNA FISH (Kubota et al., 2006b), or by alternative labeling (Coleman et al., 2007). Target functional genes. For this, there are two categories: (1) PCR-generated probes like polynucleotide

FISH (Moraru et al., 2010) or (2) amplification of the target sequence like in situ PCR (Hodson et al., 1995; Rodríguez-Castaño, G., 2008; Tani et al., 1998), in situ loop-mediated isothermal amplification (Maruyama et al., 2003); recognition of individual genes [RING]FISH (Zwirglmaier et al., 2004b; Pratscher et al., 2009; Hauer et al., in preparation), in situ rolling-circle amplification (Maruyama et al., 2005; Hoshino et al., 2010), cycling primed in situ amplification (CPRINS, Kenzaka, 2005; Tamaki et al., 2005), circularizable probes (Zhang et al., 2006), circularizable probes (Zhang et al., 2006; Hoshino et al., 2010), peptide nucleic acid assisted rolling circle amplification FISH (Smolina et al., 2007), and polynucleotide FISH (Moraru et al., 2010). Category III – combining FISH with advanced cameras, microscopes, or other analytical tools:  CCD camera (Amann and Schleifer, 2001); confocal

    

laser scanning microscopy (Wagner et al., 1998), and advanced digital image processing (Daims, 2009; Daims and Wagner, 2007) Electron microscopy (Kenzaka et al., 2005; Ishidoshiro et al., 2005) Spectral imaging (Ainsworth et al., 2006) RAMAN spectroscopy (Wagner, 2009) nanoSIMS (Behrens et al., 2008; Orphan, 2009) Atomic force microscopy (Huang et al., 2009)

Category IV – enrichment of FISH targeted cells for subsequent analyses, and combination with high-throughput technologies:  Automatic cell sorting and enrichment techniques

based on flow cytometry (Amann and Fuchs, 2008)  Magneto-FISH (Orphan, 2009; Stoffels et al., 1999)  RING-FISH (Zwirglmaier et al., 2004a)  Whole genome amplification (Yilmaz et al., 2010b)

Many of these improvements minimize at least some of the drawbacks observed with the first generation FISH protocol; however, these new approaches do also introduce other drawbacks. For example, CARD-FISH has replaced the standard FISH protocol in several cases because it produces stronger probe signals (Pernthaler et al., 2002a, b) and just recently it was shown that it can be combined with polynucleotide probes targeting functional genes in a subsequent step (Moraru et al., 2010). Nevertheless, compared to standard oligonucleotide FISH, CARD-FISH is a rather time-consuming FISH protocol and allows only the application of one probe per hybridization experiment. Furthermore, as peroxidase can be found in nearly all organisms, there may be a risk for unspecific probe reactions. Fortunately, the peroxidase can be denatured prior to the FISH procedure, however, optimal denaturation conditions appear to be species dependent and may thus require individual optimizations (Pavlekovic et al., 2009). Similarly, the recent developments of various protocols to target non-ribosomal genes

FLUORESCENCE IN SITU HYBRIDIZATION (FISH)

like various functional genes and mRNA transcripts, or combining with various analytical tools, present different advantages as well as disadvantages. For example, considerable higher costs, advanced equipment, tedious protocols, loss of probe specificity, less flexibility in terms of combination with other probes or protocols or on various organisms, problems with adequate preservation of gene target due to high decay rate (e.g., precursors or mRNA), or unpredictable cross-reactions (e.g., when incubating a complex community with radioactive substrates or stable isotopes). However, irrespective of these drawbacks that may eventually be overcome in future, it is crucial to keep in mind that our greatest hindrance for successful, unequivocal interpretations of FISH experiments in the environment is the unknown biodiversity and multitude facets of dynamics and interactions, which paradoxically seem to grow with each advancement in the science of microbiology. Thus, it appears likely that the development of novel FISH technologies will continue in parallel with the development of novel tools and our increased understanding of microbial diversity.

Different FISH applications in geomicrobiology The fields explored in geomicrobiology are extremely diverse, ranging from the deep biosphere to the atmosphere surrounding our planet Earth, and from there to the outer space. The conditions for the different ecosystems will therefore vary tremendously so that different sampling technologies and modifications of FISH protocols must be employed in order to achieve successful FISH results. So far, FISH has not been employed in geomicrobiology as often and as successfully as in other microbiological fields like environmental biotechnology (e.g., wastewater treatment plants, Nielsen et al., 2009) or in medical sciences (e.g., Bridger et al., 2010). This is because many of the ecosystems explored in geomicrobiology are often exposed to extreme conditions and generally contain low amounts of usually novel microorganisms with slow or unknown growth rates. Furthermore, special considerations must be made in terms of interpreting results with regard to the risk of contamination of samples during sampling (e.g., during drilling experiments) and to the impact of geochemical conditions on so far published FISH protocols. Because of these obstacles, many of the recent advanced developments of FISH protocols have actually been driven by microbial ecologists and geomicrobiologists (Orphan, 2009). Below follow some examples of how FISH has been employed for some of the most widely explored ecosystems in geomicrobiology (see also images in Figure 2): Aquatic systems: In general, successful FISH experiments can be performed on aquatic systems, since at least one of the main parameters for good microbial growth and dispersion of cells and nutrients, water, is abundant. Nevertheless, several parameters may introduce several biases, e.g., low nutrient status and/or naturally occurring

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autofluorescing particles (Vesey et al., 1997). Aquatic systems with high nutrient content such as contaminated lakes or even anthropogenic systems like wastewater treatment plants will contain greater amounts of highly active bacteria than oligotrophic systems like aquifers and thus produce stronger FISH probe signals. For improved FISH on oligotrophic systems, different approaches have been undertaken: Aquifers/ground water/drinking water: These ecosystems are generally dominated by planctonic cells in rather low cell numbers due to extreme conditions like oligotrophy, different redox states (e.g., anaerobic conditions), or toxic compounds (e.g., heavy metals, chlorinated drinking water or contaminated aquifers). Due to this, the general turn-over activities and thus also the ribosome content in the cells are generally extremely low. For adequate FISH evaluation, samples often need to be concentrated (e.g., by filtration) and confounding precipitates or debris removed (e.g., by cell extraction, Caracciolo et al., 2005). Early studies reported that standard FISH protocols may work well on certain aquatic samples like lakes and bottled water (Pernthaler et al., 1998; Glöckner et al., 2000; Watanabe et al., 2000; Flies et al., 2005; Loy et al., 2005); however, more recent studies report that either rainbow FISH (Sunamura and Maruyama, 2006) or CARD-FISH are more recommendable on drinking water or groundwater (Sekar et al., 2003; Nielsen et al., 2006; Wilhartitz et al., 2007; Meisinger et al., 2010a). Ocean: Similar to the freshwater systems mentioned above, oceanic ecosystems are generally dominated by planctonic cells in rather low cell numbers. However, in contrast to the conditions in the aquifers in the subsurface, the cells in oceans are often exposed to more dynamic conditions. Despite this, the ribosome content in the cells and the turn-over activities may be rather low at least for certain species – depending on location and other conditions such as oxygen concentration, light frequencies, redox conditions, nutrient conditions, flux rates. For adequate FISH evaluation of cells, samples often need to be concentrated (e.g., by filtration) and confounding precipitates or debris must be removed (e.g., by cell extraction). Early studies demonstrated that standard FISH protocols may work well (Murray et al., 1998; Glöckner, 1999; Cottrell and Kirchman, 2000; Morris et al., 2004); however during the last decade, most studies have tried to increase probe signal intensities by modifying standard FISH protocols or applying other FISH tools like polynucleotide FISH (DeLong et al., 1999; Pernthaler et al., 2002; Preston et al., 2002) or CARD-FISH (Pernthaler et al., 2002a; b; Ishii et al., 2004; Teira et al., 2004; Herndl et al., 2005). In addition to this, various combinations with different activity targeted studies based on incubation with chloramphenicol (Ouverney and Fuhrman, 1997), radioactive substrates like thymidine (Pernthaler et al., 2002) or bromodeoxyuridine (BrdU, Pernthaler and Pernthaler, 2005), or other substrates such as amino acids

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Bact

a

Arch

Dhc

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c Fluorescence In Situ Hybridization (FISH), Figure 2 (Continued)

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d Nitrous oxide reductase nosZ

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Reductive dehalogenase RDase

10 µm

e Fluorescence In Situ Hybridization (FISH), Figure 2 Demonstration of different images from different ecosystems produced by different FISH techniques.

(Ouverney and Fuhrman, 2000; Cottrell, 2000; Cottrell, 2003), have been undertaken. Soil/sediments/subseafloor: Soil and sediments are rather difficult environments for successful FISH studies, as these habitats are extremely diverse, complex, and subject to harsh and fluctuating conditions in terms of, e.g., water content and redox conditions. Thus, more than most other ecosystems, the biosphere of microorganisms in soil is often characterized by micro-niches which may be exposed to dramatic changes with regard to space and time. In addition to this, many substances in soil or sediments produce strongly confounding background such as humic compounds, mineral precipitates or other microscopic particles of unknown character. For successful FISH experiments, two different main approaches have been made (for a review, see Schmid et al., 2006; Schmid et al., 2007): (1) Spatial separations of cells or micro-niches by approaches such as embedding and

sectioning of soil columns or by confocal laser scanning microscopy of soil aggregates, different cell extraction procedures such as density centrifugation or cation exchange membranes (McDonald, 1986; Smith and Stribley, 1994; Caracciolo et al., 2005; Kurola et al., 2005; Lunau et al., 2005; Bertaux et al., 2007), or flow cytometry (Kalyuzhnaya et al., 2006; Podar et al., 2007). (2) The second approach employed a variety of different FISH protocols. The early studies reported generally only partial success or even contradictory results between FISH and other analytical tools (Hahn et al., 1992; Ludwig et al., 1997; Zarda et al., 1997; Chatzinotas, 1998; LlobetBrossa et al., 1998; Christensen, 1999; Hristova et al., 2000; Ravenschlag et al., 2000; Janvier et al., 2003; Kobabe et al., 2004; Rusch and Amend, 2004; Flies et al., 2005; Knittel et al., 2005). However, during the last years, most studies have now replaced the standard FISH tool with CARD-FISH and nanoSIMS (Ishii et al., 2004;

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Mussmann et al., 2005; Schippers et al., 2005; Tal et al., 2005; Orphan 2009). Karst regions/rock/deep biosphere: During the last decade, our understanding of the role of microorganisms and their interactions with rocks and minerals over geological timescales has expanded considerably. The interaction of minerals with water and microbes do not only influence soil fertility, transport compounds, and pollutants through the subsurface, but they also contribute to dissolution of rocks or precipitation of various mineral formations like speleothems in caves and may thus even play a key role in global climate change (Northup and Lavoie, 2001; Brown and Lee, 2007; Cockell and Herrera, 2008). Several attempts have been made to screen for microbial life in rocks and set up validated guidelines for this (Barton et al., 2001; Barton et al., 2006; Tobin et al., 1999). The reoccurring concern is to not only prove extant microbial processes (Orphan and House, 2009) but also current cellular-based in situ activities in often carbonpoor systems. Since it may be difficult to retrieve sufficient amounts of RNA or other activity biomarkers from these systems due to low cell counts and activities, FISH, in particular in combination with advanced tools like nanoSIMS, may play a crucial role for demonstrating whole cells with targetable genes (Orphan, 2009). Despite these obstacles, standard FISH as well as CARD-FISH protocols have been successfully applied on, e.g., microbial mats in association with water and energy-rich substances for life based on chemolithoautotrophy in rocky environments or caves (Engel et al., 2003; Macalady et al., 2006; Meisinger et al., 2007; Meisinger et al., 2010b). However, certain types of microbial mats may be strongly hampered by confounding background caused by various mineral precipitates (Engel et al., 2010; Koebberich, 2008). Rigid environments like calcified biofilms or speleothems thus demand sophisticated modifications of standard FISH protocols as well as CARD-FISH (e.g., Shiraishi et al., 2008). Extremophiles in various geological environments: Extremophiles thrive in extreme ecosystems which are characterized by extreme physical or geochemical conditions. Such conditions may pose a particular challenge for FISH experiments. So far, FISH has only been practiced in certain types of extremophilic ecosystems with mixed success, e.g., in hot springs (Nubel et al., 2002; Simbahan et al., 2005; Nakagawa et al., 2006; Weidler et al., 2008), polar regions (Junge et al., 2004), halophilic systems (Rossello-Mora et al., 2003; Maturrano et al., 2006), mines/bioleaching systems (Schrenk et al., 1998; Edwards et al., 1999; Bond et al., 2000; Takai, 2002; Baker et al., 2004), hydrothermal vents (Harmsen et al., 1997; Schrenk et al., 2003; Nakagawa et al., 2006), and methane hydrate seeps (Boetius and al, 2000).

Summary FISH (fluorescence in situ hybridization) is a valuable tool for in situ identification of microbial community structure, cell morphology, dynamics and spatial distribution.

If combined with different analytical tools additional information of activity, function and interactions of different species on single cell level can be performed. However, for successful FISH experiments, it is crucial to optimize and validate all steps throughout the FISH protocol, ranging from proper sampling technique and fixation, selection of appropriate gene markers, hybridization conditions and relevant controls, to optimal detection and evaluation strategies.

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Cross-references Anaerobic Oxidation of Methane with Sulfate Basalt (Glass, Endoliths) Biogeochemical Cycles Biomarkers (Molecular Fossils) Immunolocalization Isotopes and Geobiology Karst Ecosystems Metagenomics Microbial Communities, Structure, and Function

FORAMINIFERA

Microbial Ecology of Submarine Caves Protozoa (Heterotroph, Eukaryotic) RNA World Scanning Probe Microscopy Speleothems Terrestrial Deep Biosphere

FORAMINIFERA Alexander V. Altenbach Ludwig-Maximilians-University Munich, Munich, Germany

Synonyms Forams (for short) Definition Foraminifera (from ancient Greek “hole bearers”, refers to the pores that early observers recognized in the calcitic tests) are amoeboid, eukaryotic protists with a large network of very thin cytoplasm extrusions (reticulopodia). Most species produce a genetically fixed shell (test), made of organic matter, agglutinated particles, calcite, aragonite, high-magnesium calcite, or opaline silica. The tests may be either simple bowls or tubes, or branched, coiled, substructured, and ornamented in diverse complexity. Several thousand species are found living in modern oceans, and this number still increases. Several times more extinct species are recovered from the fossil record. Foraminifera are considered an order (sometimes a class) in paleontology, subdivided by the structure and composition of modern and fossil tests. They are ranked among the “Granuloreticulosa” in biology, characterized by (1) their branching and anastomosing networks of reticulopodia, (2) a rapid bidirectional transport of intracellular granules and membrane domains driven by specific tubulin assemblies, and (3) a complex alternation of sexual and asexual reproductive cycles. Genetic investigations led to the definition of the first rank group “Foraminifera” within the Supergroup “Rhizaria”. Independent from such rankings, they range among the most common, diverse, and ubiquitous groups of single celled, test building eukaryotes on earth. Introduction Numerous foraminiferal species indicate specific environmental conditions, and the first and last appearance of extinct species exactly indicates time intervals in earth history. Investigations on the chemical and isotopical composition of their calcitic tests reveal environmental drifts in global ice volumes, palaeo-temperatures and -salinities of marine water masses, and marine palaeo-productivity (Hemleben et al., 1989; Murray, 2006). Despite the long term and extensive attention on foraminifera in

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taxonomy (Loeblich and Tappan, 1988) and environmental sciences (Sen Gupta, 1999), our comprehensions on their evolution and ecophysiology are still in transition. Molecular ecology and metagenomics applied to the study of microbial biodiversity are changing our view of the biosphere. An impressive extension of foraminiferal clades has been uncovered by such molecular tools: naked freshwater amoebians, naked marine giant amoebians, and giant deep-sea protists formerly considered of unknown origin, where added to the first rank group Foraminifera (Adl et al., 2005; Pawlowski and Burki, 2009). Even amoeboid parasites of terrestrial plants are supposed to reach out from this lineage (Archibald and Keeling, 2004). Efforts to couple the growing phylogenetic diversity of single-celled organisms with ecological function lead to the discovery of novel metabolisms, processes, and symbiosis, and thus to a reevaluation of the geobiochemical impact of pro- and eukaryotes (Moya et al., 2008).

Earth history The earliest fossils presumed for the foraminiferal lineage reach back into the Mesoproterozoic, despite severe problems in defining even Ediacaran remainders (Dong et al., 2008). The Cambrian provides already several foraminiferan families with organic-walled or agglutinated tests (Benton, 1993). A significant eukaryote radiation occurs in the Ordovician, coincident with increasing nutrient supply, enhanced primary production, and the propagation of planktonic metazoan stages. Such amplified planktonic food chains sponsored increased flux rates of organic matter at the sea floor, and thus the radiation of planktotrophy (Nützel et al., 2006). The calcitic fusulinids start in the Silurian (Loeblich and Tappan, 1988), and extend into the first rhizarian, rock forming mass deposition of biogenic shallow water carbonates, from the Pennsylvanian to the Permian. The fusulinid clade develops all principal morphological adaptations known from modern benthic foraminifera sheltering phototrophic symbionts, thus these mass deposits should result from host–symbiont interactions (Anderson and Lee, 1991). The structure of modern marine primary production was based on the evolution of diatoms, coccoliths, and modern dinoflagellates in the Triassic to Jurassic (Falkowski et al., 2004). Restructuring and enhancing marine primary productivity to an elevated level is recorded by the boost of rotaliid foraminifera, modern metazoans, and increasing bioturbation (Martin et al., 2008). One tribe of calcitic foraminifera, the globigerinids, completely adapted to this new planktonic food chain, and started to enter a purely pelagic live style in the Middle Jurassic. The boost of rock forming biogenic carbonate (chalk) and silica (diatomite) production from these new planktonic systems, however, was delayed to the Cretaceous in the marine realm, and to the Eocene for lacustrine diatomites. The first major radiation of planktonic foraminifera occurred during the mid-Cretaceous. The density structure of the upper water

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column and the formation of deeper water masses were altered by plate tectonic activities, a major long-term rise of global sea level, and an overall increase of global temperatures (Leckie, 1989; Martin et al., 2008). Following the massive mass extinction of planktonic foraminifera at the K/T boundary, symbiosis with phototrophic endosymbionts was one key factor for the radiation of Cenozoic lineages (Norris, 1996). The diversity of benthic foraminifera, and thus presumably their ecological demands and adaptations, were influenced more modestly. The first appearance of foraminiferal tests build up by opaline silica occurs in the Miocene, by the benthic foraminiferal suborder Silicoloculinina (Loeblich and Tappan, 1988). Enhanced terrestrial silica weathering and riverine silica input by grassland expansion offers a causally driven correlation, similar to marine diatom diversification (Falkowski et al., 2004).

Modern oceans Planktonic foraminifera are most influential for the open marine calcite budget. Their annual flux rate of calcitic tests at 100 m water depth accounts for 1.3–3.2 Gt, equivalent to 23–56% of the global, open marine CaCO3 flux. During most of the year, dissolution takes place within the upper water masses (>700 m water depth), and only 1–3% of the initially exported CaCO3 reaches the seafloor. This biogeochemical pump is most influential for the alkalinity of surface and intermediate water masses. During short pulsed flux events, mass dumps of fast settling particles yield the major contribution to the formation of deep sea sediments. This pathway transfers 0.4–0.9 Gt CaCO3 annually, accounting for 32–80% of the total deep-marine calcite budget (Schiebel, 2002). Maximum carbonate production appears in open ocean realms where symbiotic and non-symbiotic planktonic foraminifera occur syntopic (Zaric et al., 2006). The contribution of coccolithophores (mean 12%) and calcareous dinophytes (0–3.5%) is comparably small (Schiebel, 2002; Sarmiento et al., 2002). Planktic gastropods (pteropods) may add another 10% only locally (Schiebel, 2002). Phototrophic endosymbionts play a major role in the carbonate production of planktic foraminifera. Such endosymbionts are the key factor for the foraminiferal successes in nutrient depleted environments, and for their enhanced turnover (Hemleben et al., 1989; Lee, 2006). One sixth of the global carbonate production is shared by reef complexes, dominated by symbiont-bearing corals, foraminifera, and coralline algae. With an annual production of 130 million tons of CaCO3, benthic foraminifera bearing phototrophic symbionts contribute nearly 5% of the annual present-day carbonate production in the world’s reef and upper shelf areas (Langer, 2008). Marine benthic foraminifera without phototrophic symbionts are much more diverse, inhabiting all ocean realms from estuaries to deep ocean trenches. But their share in marine biogenic carbonate production ranges only near 1% (Langer, 2008). This is due to the rapidly decreasing

flux rate and nutritional value of organic matter reaching the deeper ocean floors (Altenbach and Struck, 2001). Deep-sea foraminifera have a metabolic turnover comparable to bacteria (Linke, 1992; Moodley et al., 2002), and their grazing rate exceeds that of common meiofaunal groups (Pascal et al., 2008). They have a considerable share in abyssal respiration rates (Witte et al., 2003), but this does not sum up to a significant impact in biogenic carbonate production. Complete denitrification was observed as an endogenic metabolic pathway for benthic foraminifera in laboratory cultures (Risgaard-Petersen et al., 2006). The species facultatively switch to denitrification when high loads of organic matter are available in oxygen-depleted environments. Field observations in the upwelling region off Chile revealed that benthic denitrification is dominated by foraminifera, and not by bacteria (Høgslund et al., 2008). In view of the large benthic foraminiferal populations recovered from other oxygen minimum zones, and the large uncertainties in the global nitrogen budget (Gruber and Galloway, 2008), these findings may challenge our considerations on marine denitrification cycles (Høgslund et al., 2008). Several species related to denitrification acquire chloroplasts from settled algae in the deep ocean, which are kept functional in the foraminifers cytoplasm for at least one year (Grzymski et al., 2002). Such kleptoplasts produce hydroxyl radicals, which offer an alternative source of oxygen for the hosts metabolism by H2O2 breakdown (Bernhard and Bowser, 2008). Sulfur-oxidizing bacterial endosymbionts were considered for benthic foraminifera recovered from longterm sulfidic environments in the deep sea (Bernhard et al., 2006) and from temporal anoxic shelf areas (Bernhard, 2003). However, the definition of oxic or anoxic conditions is a problem by itself near the redox cline, as the interwoven, submillimetric structures of differing redox conditions build up a foamy, three dimensional space. Contrasting microbial turnover takes place in smallest pore water compartments. The interlacement may be so dense that foraminiferal inhabitants can not be attributed to either oxic or anoxic conditions (Bernhard et al., 2003). Xenophyophores and Komokiaceans are very common deep-sea foraminifera with a comparatively large and loosely agglutinated test. They may cover up to 50% of hadal seafloors (Lemche et al., 1976). Early research on the role of Xenophyophores and Komokiaceans for biogenic metal dissolution and precipitation considered a significant impact for the formation of manganese nodules (Riemann, 1985). The microscopic barite crystals, commonly found in the endoplasm of Xenophyophores (Gooday and Nott, 1982), were considered a proof for their endogenic barite precipitation (Bertram and Cowen, 1997). However, these rhizarians ingest considerable amounts of highly refractory organic matter and produce large amounts of waste pellets (stercomata), offering an attractive food source for deep ocean microbial communities (Nozawa et al., 2006). Gardening of a specialized

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bacterial flora growing on the stercomes of Xenophyophores was made plausible by Laureillard et al. (2004). Such affiliated microbial communities will move the redox state by degrading the stercomata, a factor that should interplay with the observed metal reactions. Gathering refractory material that is transferred to higher levels of the food chain by bacterial counterparts is highly effective. Parallel to the increasing number of bacteria available as food, it can enhance the metabolic pathways of the host. Limited endogenous proteolytic capacities of benthic organisms are surpassed by enzyme sharing with bacteria, in most cases combined with the production of an agglutinated detrital lump or burrow (Riemann and Helmke, 2002). Benthic foraminifera from the earliest tribes to most recent lineages retain the ability to rapidly produce such detrital lumps or cysts, in addition to the genetically fixed test (Heinz et al., 2005). Most researchers make reasonable tropical adaptations and bacterial gardening for the common production of such cysts (Heinz et al., 2005).

Conclusions The general outline of foraminiferal ecology and their overall impact on the marine food chains and calcite budgets is well documented and understood. Recently discovered, endogenic denitrification offers a facultative metabolic pathway for benthic foraminifera with potential impact for the marine nitrogen cycle. Additional complexity in synecology and ecophysiology is also indicated by observations on chemotrophic bacterial endosymbionts and bacterial gardening, not fully understood at present. Chemotrophic symbionts and functional kleptoplasts seem to play a key role for anoxic and sulfidic microenvironments. We may presume that foraminiferal interactions with bacteria play a more significant role in geobiology and biogeochemical cycles than has previously been suspected. Bibliography Adl, S. M., Simpson, A. G. B., Farmer, M. A., Andersen, R. A., Anderson, O. R., Barta, J. R., Bowser, S. S., Brugerolle, G., Fensome, R. A., Fredericq, S., James, T. Y., Karpov, S., Kugrens, P., Krug, J., Lane, C. E., Lewis, L. A., Lodge, J., Lynn, D. H., Mann, D. G., Mccourt, R. M., Mendoza, L., Moestrup, Ø., Mozley-Standridge, S. E., Nerad, T. A., Shearer, C. A., Smirnov, A. V., Spiegel, F. W., and Taylor, M. F. J. R., 2005. The new higher level classification of eukaryotes with emphasis on the taxonomy of protists. The Journal of Eukaryotic Microbiology, 52, 399–451. Altenbach, A. V., and Struck, U., 2001. On the coherence of organic carbon flux and benthic foraminiferal biomass. Journal of Foraminiferal Research, 31, 79–85. Anderson, O. R., and Lee, J. J., 1991. Symbiosis in foraminifera. In Lee, J. J., and Anderson, O. R. (eds.), Biology of Foraminifera. London: Academic Press, pp. 157–220. Archibald, J. M., and Keeling, P. J., 2004. Actin and ubiquitin protein sequences support a Cercocoan/Foraminiferan ancestry for the Plasmodiophorid plant pathogenes. Journal of Eukaryotic Microbiology, 51, 113–118.

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Benton, M. J., 1993. The Fossil Record 2. London: Chapmann & Hall. Bernhard, J. M., 2003. Potential symbionts in bathyal foraminifera. Science, 299, 861. Bernhard, J. M., and Bowser, S. S., 2008. Peroxisome proliferation in foraminifera inhabiting the chemocline: An adaptation to reactive oxygen species exposure? Journal of Eukaryotic Microbiology, 55, 135–144. Bernhard, J. M., Visscher, P. T., and Bowser, S. S., 2003. Submillimeter life positions of bacteria, protists, and metazoans in laminated sediments of the Santa Barbara Basin. Limnology Oceanography, 48, 813–828. Bernhard, J. M., Habura, A., and Bowser, S. S., 2006. An endobiontbearing allogromiid from the Santa Barbara Basin: implications for the early diversification of foraminifera. Journal of Geophysical Research, 111, G03002, DOI:10.1029/2005JG000158. Bertram, M. A., and Cowen, J. P., 1997. Morphological and compositional evidence for biotic precipitation of marine barite. Journal of Marine Research, 55, 577–593. Dong, L., Xiao, S., Shen, B., and Zhou, C., 2008. Silicified Horodyskia and Palaeopascichnus from upper Ediacaran cherts in South China: tentative phylogenetic interpretation and implications for evolutionary stasis. Journal of the Geological Society U. K., 165, 367–378. Falkowski, P. G., Katz, M. E., Knoll, A. H., Quigg, A., Raven, J. A., Schofield, O., and Taylor, F. J. R., 2004. The evolution of modern eukaryotic phytoplankton. Science, 305, 354–360. Gooday, A. J., and Nott, A. J., 1982. Intracellular barite crystals in two Xenophyophores, Aschemonella ramuliformis and Galatheammina sp. (Protozoa, Rhiziopoda) with comments on the taxonomy of A. ramuliformis. Journal of the Marine Biological Association U. K., 62, 595–605. Gruber, N., and Galloway, J. N., 2008. An Earth-system perspective of the global nitrogen cycle. Nature, 451, 293–296. Grzymski, J., Schofield, O. M., Falkowski, P. G., and Bernhard, J. M., 2002. The function of plastids in the deepsea benthic foraminifer, Nonionella stella. Limnology and Oceanography, 47, 1569–1580. Heinz, P., Geslin, E., and Hemleben, C., 2005. Laboratory observations of benthic foraminiferal cysts. Marine Biology Research, 1, 149–159. Hemleben, C., Spindler, M., and Anderson, O. R., 1989. Modern Planktonic Foraminifera. New York: Springer. Høgslund, S., Revsbech, N., Cedhagen, T., Nielsen, L., and Gallardo, V., 2008. Denitrification, nitrate turnover, and aerobic respiration by benthic foraminiferans in the oxygen minimum zone off Chile. Journal of Experimental Marine Biology and Ecology, 359, 85–91. Langer, M. R., 2008. Assessing the contribution of foraminiferan protists to global ocean carbonate production. Journal of Eukaryotic Microbiology, 55, 163–169. Laureillard, J., Mejanelle, L., and Sibuet, M., 2004. Use of lipids to study the trophic ecology of deep-sea xenophyophores. Marine Ecology Progress Series, 270, 129–140. Leckie, R. M., 1989. A paleoceanographic model for the early evolution history of planktonic foraminifera. Palaeogeography, Palaeoclimatology, Palaeoecology, 73, 107–138. Lee, J. J., 2006. Algal symbiosis in larger Foraminifera. Symbiosis, 42, 63–75. Lemche, H., Hansen, B., Madsen, E. J., Tendal, O. S., and Wolf, T., 1976. Hadal life as analyzed from photographs. Videnskabelige meddelelser fra Dansk naturhistorisk forening, 139, 263–336. Linke, P., 1992. Metabolic adaptions of deep-sea benthic foraminifera to seasonally varying food input. Marine Ecology Progress Series, 81, 51–63.

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Loeblich, A. R., and Tappan, H., 1988. Foraminiferal Genera and their Classification. New York: Van Nostrand Reinhold. [year of publication often cited erroneously as 1987; see Loeblich, A. R., and Tappan, H., 1989. Publication date of foraminiferal genera and their classification. Journal of Paleontology, 63, 253.] Martin, R. E., Quigg, A., and Podkovyrov, V., 2008. Marine Biodiversification in response to evolving phytoplankton stoichiometry. Palaeogeography Palaeoclimatology Palaeoecology, 258, 277–291. Moodley, L., Middelburg, J. J., Boschker, H. T. S., Duineveld, G. C. A., Pel, R., Herman, P. M. J., and Heip, C. H. R., 2002. Bacteria and foraminifera: key players in a short-term deep-sea response to phytodetritus. Marine Ecology Progress Series, 236, 23–29. Moya, A., Pereto, J., Gil, R., and Latorre, A., 2008. Learning how to live together: genomic insight into prokaryote-animal symbiosis. Nature Reviews Genetics, 9, 218–229. Murray, J. W., 2006. Ecology and Applications of Benthic Foraminifera. Cambridge: Cambridge University Press. Norris, R. D., 1996. Symbiosis as an evolutionary innovation in the radiation of Paleocene planktic foraminifera. Paleobiology, 22, 461–480. Nozawa, F., Kitazato, H., Tsuchiya, M., and Gooday, A. J., 2006. ‘Live’ benthic foraminifera at a abyssal site in the equatorial Pacific nodule province: abundance, diversity and taxonomic composition. Deep-Sea Research I, 53, 1406–1422. Nützel, A., Lehnert, O., and Fryda, J., 2006. Origin of planktotrophy - evidence from early molluscs. Evolution and Development, 8, 325–330. Pascal, P., Dupuy, C., Mallet, C., Richard, P., and Niqul, N., 2008. Bacterivory in benthic organisms in sediment: quantification using 15N-enriched bacteria. Journal of Experimental Biology and Ecology, 355, 18–26. Pawlowski, J., and Burki, F., 2009. Untangling the phylogeny of amoeboid protists. Journal of Eukaryotic Microbiology, 56(1), 16–25. Riemann, F., 1985. Eisen und Mangan in pazifischen TiefseeRhizopoden und Beziehungen zur Manganknollen-Genese. Internationale Revue der gesamten Hydrobiologie, 70(1), 165–172. Riemann, F., and Helmke, E., 2002. Symbiotic relations of sediment-agglutinating nematodes and bacteria in detrital habitats: the enzyme sharing concept. Marine Ecology, 23, 93–113. Risgaard-Petersen, N., Langezaal, A. M., Ingvardsen, S., Schmid, M. C., Jetten, M. S. M., Op den Camp, H. J. M., Derksen, J. W. M., Pina-Ochoa, E., Eriksson, S. P., Nielsen, L. P., Revsbech, N. P., Cedhagen, T., and van der Zwaan, G. J., 2006. Evidence for complete denitrification in a benthic foraminifer. Nature, 443, 93–96. Sarmiento, J. L., Dunne, J., Gnanadesikan, A., Key, R. M., Matsumoto, K., and Slater, R., 2002. A new estimate of the CaCO3 to organic carbon export ratio. Global Biogeochemical Cycles, 16(4), no 1107. doi:10.1029/2002/GB001919. Schiebel, R., 2002. Planktic foraminiferal sedimentation and the marine calcite budget. Global Biogeochemical Cycles, 16(4), no 1065, doi:10.1029/2001GB001459. Sen Gupta, B., 1999. Modern Foraminifera. Dordrecht: Kluwer Academic Press. Witte, U., Wenzhöfer, F., Sommer, S., Boetius, A., Heinz, P. N. A., Sand, M., Cremer, A., Abraham, W. B. J. B., and Olaf, P., 2003. In situ experimental evidence of the fate of a phytodetritus pulse at the abyssal sea floor. Nature, 424, 763–766. Zaric, S., Schulz, M., and Mulitza, S., 2006. Global prediction of planktonic foraminiferal fluxes from hydrographic and productivity data. Biogeosciences, 3, 187–207.

Cross-references Algae (Eukaryotic) Bioerosion Carbonate Environments Carbonates Chemolithotrophy Divalent Earth Alkaline Cations in Seawater Isotopes and Geobiology Protozoa (Heterotroph, Eukaryotic) Symbiosis

FRUTEXITES Marta Rodríguez-Martínez1, Christine Heim2, Nadia-Valérie Quéric2, Joachim Reitner2 1 Universidad de Alcalá, Alcalá de Henares, Madrid, Spain 2 University of Göttingen, Göttingen, Germany

Synonyms Colloform limonitic crusts; Frutexites crusts; Frutexiteslike forms; Frutexites microstromatolite; Frutexites tuffs; Haematitic/ferruginous/iron microstromatolites; Iron dendritic aggregates; Iron shrubs; Pillar-shaped microstromatolites

Frutexites, Figure 1 Original figures of genus of Frutexites described by Maslov (1960).

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Definition Frutexites is a problematic microfossil rich in iron. From a taxonomic point of view, only five species have been figured (Frutexites arboriformis, Maslov, 1960; F. microstroma, Walter and Awramik, 1979; Frutexites sp. 1, Frutexites sp. 2, and New gen. 3, Tsien, 1979), although the authors mostly use the term Frutexites sensu lato. The genus Frutexites was coined by Maslov (1960) in order to describe submillimeter-sized, iron-rich, and subordinate calcite microfossils (Figure 1). Frutexites have a dendritic shape formed by divergently branched microcolumns. The height and width of microcolumns as well as their composition and microstructure can vary (Table 1). The preservation of microstructure is strongly controlled by its dominant mono- or polymineral

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character. Microstructure is formed by convex-upward laminae, which sometimes show radially arranged fibers. Frutexites can occur mainly as monomineral as well as polymineral structures with: iron-rich and/or manganeserich (hematites, iron hydroxides, Fe-Mn oxyhydroxides), and/or carbonate-rich (calcite, ferroan calcite, dolomite), and/or siliciclastic-rich (argillite, microquartz), and/or phosphatic-rich zones.

Depositional environmental occurrences Frutexites has been described in marine environments such as shallow and deep water stromatolites, microbial limestones, hardgrounds, and condensed pelagic limestones as well as in cavities, sheet-cracks, veins, and Neptunian dikes. However, comparisons with

Frutexites, Table 1 Sizes, morphological and compositional parameters of Frutexites according to different authors Authors

Height

Maslov (1960)

Up to 400 µm 25–30 µm, (see more of Figure 1) 50 µm 20–500 µm 10–200 µm

Horodyski (1975) Walter and Awramik (1979) Myrow and Coniglio (1991) Böhm and Brachert (1993) Woods and Baud (2008)

Width

Microstructure

Morphology

Composition

Sheets with occasionally circular spaces

Radially diverging and branched sheets

Iron hydroxide and carbonate

2–10 µm convex-upward Pillar-shaped branched and not laminae Up to 450 µm 5–120 µm 0.7–2.7 µm convex upward Undulose layers, laminae with laminae and axial tube protruding pustules and erect (trichome?) branching microcolumns Unbranching and branching columns 250 µm–4 75–600 µm, Chambers, laminae, fibers, mm, most 250 µm and projections 200 mg). The impact of the use of microbial activity for the solution and precipitation of gold was undervalued for a long period, thus affording the opportunity for economic and environmentally friendly gold bio-processing. Furthermore, the current high price of gold has led to intensive research into the precipitation of gold. Some iron and sulfur oxidizers (e.g., Acidothiobacillus ferrooxidans, Acidithiobacillus thiooxidans) contribute to the disintegration of important gold ores (Nordstrom and Southam, 1997). In an environment of gold-bearing rocks, the dissolved gold content is 40 times higher than in comparable non-gold-bearing areas (Benedetti and Boulègue,

0.3 mm

Gold, Figure 1 A secondary electron (SE) image of a hexagonal gold crystal from the secondary gold deposit Silberkuhle near Korbach, Germany.

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10 µm

Gold, Figure 2 Laser scanning micrograph (ex. 488 nm, em. 543 nm) of microorganisms (green) on a gold grain (red) from the Eder River, Germany. Eubacteria labeled by oligonucleotide probe EuB-CY3.

and not in the surrounding sediments. With a similarity of 99%, the sequence correlated to Cupriavidus (Ralstonia) metallidurans. Ninety percent of the bacteria in the laboratory experiment died because of the toxicity of AuCl
4. The surviving cells adapted themselves to the high concentration of Au(III) ions and precipitated gold. In addition to morphological studies (Bischoff, 1997), this provides evidence for bioaccumulation of secondary gold (e.g., nuggets, gold grains) by bacteria like C. metallidurans (Figure 2). The basal biochemical microbial processes of the biomineralization of gold are not completely understood yet. Precipitation of gold provides the cellular defense of C.metallidurans and is controlled by a coupling of efflux, reduction and possibly methylation of gold complexes (Reith et al., 2009). Reith et al. (2007) have published a comprehensive review of the geomicrobiology of gold.

Summary Different microorganisms are partially resistant to dissolved gold. Various microbial activities precipitate gold. Biofilms of Cupriavidus metallidurans accumulate gold naturally and contribute to the formation of secondary gold. Bibliography Benedetti, M., and Boulègue, J., 1991. Mechanism of gold transfer and deposition in a supergene environment. Geochimica et Cosmochimica Acta, 55, 1539–1547.

Bischoff, G. C. O., 1997. The biological origin of bacterioform gold from Australia. Neues Jahrbuch Geologische Paläontologische Abhandlungen, H6, 329–338. Karthikeyan, S., and Beveridge, T. J., 2002. Pseudomonas aeruginosa biofilms react with and precipitate toxic soluble gold. Environmental Microbiology, 4(11), 667–675. Kashefi, K., Tor, J. M., Nevin, K. P., and Lovley, D. R., 2001. Reductive precipitation of gold by dissimilatory Fe(III)-reducing bacteria and archaea. Applied and Environmental Microbiology, 67(7), 3275–3279. Lengke, M. F., and Southam, G., 2005. The effect of thiosulfateoxidizing bacteria on the stability of the gold–thiosulfate complex. Geochimica et Cosmochimica Acta, 69(15), 3759–3772. Lengke, M. F., Ravel, B., Fleet, M. E., Wanger, G., and Southam, G., 2006. Mechanisms of gold bioaccumulation by filamentous cyanobacteria from gold(III)–chloride complex. Environmental Science & Technology, 40(20): 6304–6309. Nakajima, A., 2003. Accumulation of gold by microorganisms. World Journal of Microbiology, 19, 369–374. Nordstrom, D. K., and Southam G., 1997. Geomicrobiology of sulfide mineral oxidation. Reviews in Mineralogy and Geochemistry, 35, 361–390. Reith, F., Rogers, S., McPhail, D. C., and Webb, D., 2006. Biomineralization of gold: biofilms on bacterioform gold. Science, 313, 333–336. Reith, F., Lengke, M. F., Falconer, D., Craw, D., and Southam, G., 2007. The geomicrobiology of gold. ISME Journal, 1, 567–584. Reith, F., Etschmann, B., Grosse, C., Moors, H., Benotmane, M. A., Monsieurs, P., Grass, G., Doonan, C., Vogt, S., Lai, B., MartinezCriado, G., George, G. N., Nies, D. H., Mergeay, M., Pring, A., Southam, G., and Brugger, J., 2009. Mechanisms of gold biomineralization in the bacterium Cupriavidus metallidurans. Proceedings of the National Academy of Sciences of the United States of America, 106(1), 17757–17762. Sand, W., Gehrke, T., Jozsa, P.-G., and Schippers, A., 2001. (Bio) chemistry of bacterial leaching – direct vs. indirect bioleaching. Hydrometallurgy, 59, 159–175. Southam, G., and Beveridge, T. J., 1996. The occurrence of sulfur and phosphorus within bacterially derived crystalline and pseudocrystalline octahedral gold formed in vitro. Geochimica et Cosmochimica Acta, 60(22), 4369–4376.

Cross-references Algae (Eukaryotic) Archaea Bacteria Biofilms Fungi and Lichens Microbial Biomineralization Ores, Microbial Precipitation and Oxidation

GONDWANALAND, FORMATION Joseph G. Meert University of Florida, Gainesville, FL, USA

Definition Gondwanaland or “Gondwana” is the name for the southern half of the Pangaean supercontinent that existed some 300 million years ago. Gondwanaland is composed of the major continental blocks of South America, Africa,

GONDWANALAND, FORMATION

Arabia, Madagascar, Sri Lanka, India, Antarctica, and Australia (Figure 1). The name “Gondwana” is derived from a tribe in India (Gonds) and “wana” meaning “land of.” Gondwanaland is superficially divided into a western half (Africa and South America) and an eastern half (India, Sri Lanka, Madagascar, Antarctica, and Australia). The archetypal view of Gondwanaland assembly was an Ediacaran–Cambrian-age coalescence of East Gondwana (India, Sri Lanka, Madagascar, Australia, and Antarctica) with West Gondwana (South America and Africa) along the Mozambique Belt (labeled East African Orogen in Figure 1). “Pan-African” tectonothermal belts outside of the Mozambique Belt (500–600 Ma) were well-known at the time, but they were often treated as zones of ensialic activity rather than sites of continental collision. Although this rather simplistic view of Gondwanaland assembly is now strongly debated, the polyphase assembly model outlined below is not completely accepted (see Yoshida, 2007). A new view of Gondwanaland assembly began with the work by Stern (1994) who demonstrated clear evidence of juvenile arc development and accretion in the Arabian– Nubian shield and continental collision to the south in Kenya and Tanzania. In the early 1990s, work by Dalziel (1991), Hoffman (1991), and Moores (1991) hinted that

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the assembly of Gondwanaland followed the breakup of an earlier supercontinent called Rodinia (see Chapter Breakup of Rodinia). Although the exact continental configuration of Rodinia is still debated (for example, Meert and Torsvik, 2003), the assembly of West Gondwana is more or less viewed as a polyphase accretion of formerly disparate blocks while East Gondwana is generally treated as a coherent unit from 1,100 Ma until its breakup in the Mesozoic (Meert, 2003). More recently, Fitzsimons (2000), Meert (2003), Collins and Pisarevsky (2005), and Kelsey et al. (2008) noted that at least two “Pan-African” age mountain belts cut across the East Antarctic shield and juxtapose distinct crustal fragments thought to comprise the East Antarctic “Grenvillian craton.” The existence of these belts preclude a united East Gondwana and favor a polyphase accretion of major blocks to form Greater Gondwanaland in the Cambrian. In this scenario, the assembly of Gondwanaland was accomplished along three major orogenic belts known as the Brasiliano, East African, and Kuungan orogenies (Figure 1). Additional data supporting a polyphase assembly of East Gondwana are derived from paleomagnetic and geochronologic studies. No matter what the exact model of Gondwanaland assembly is, the formation of this supercontinent followed

Gondwanaland, Formation, Figure 1 The Gondwanaland supercontinent. The cratons comprising West Gondwana are shaded in light blue and those comprising East Gondwana are shaded in yellow. Neoproterozoic orogenic belts crisscross the supercontinent. Those associated with the final amalgamation of the supercontinent are the East African Orogen (750–620 Ma; red), the Brasiliano-Damara Orogen (630–520 Ma; blue), and the Kuungan Orogen (570–530 Ma; green).

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shortly after the extreme glaciations during the late Neoproterozoic (Hoffman et al., 1998) and was nearly synchronous with the rapid evolutionary pulse that coincides with the Ediacaran and Cambrian radiations. Final assembly of Gondwanaland and a circumGondwanaland mountain chain is thought to have provided nutrients to the oceanic realm and also resulted in a large 87Sr/86Sr spike due to the erosion of the mountains (Squire et al., 2006). Therefore, links between the assembly of the Gondwanaland supercontinent and biological evolution are forwarded by numerous studies (Meert and Lieberman, 2008).

Bibliography Collins, A. S., and Pisarevsky, S. A., 2005. Amalgamating eastern Gondwana: the evolution of the circum-Indian orogens. Earth Science Reviews, 71, 229–270. Dalziel, I. W. D., 1991. Pacific margins of Laurentia and East Antarctica as a conjugate rift pair: evidence and implications for an Eocambrian supercontinent. Geology, 19, 598–601. Fitzsimons, I. C. W., 2000. A review of tectonic events in the East Antarctic Shield, and their implications for three separate collisional orogens. Journal of African Earth Sciences, 31, 3–23. Hoffman, P. F., 1991. Did the breakout of Laurentia turn Gondwanaland inside out? Science, 252, 1409–1412. Hoffman, P. F., Kaufman, A. J., Halverson, G. P., and Schrag, D. P., 1998. A Neoproterozoic snowball Earth. Science, 281, 1342–1346. Kelsey, D. E., Wade, D. P., Collins, A. S., Hand, M., Sealing, C. R., and Netting, A., 2008. Discovery of a Neoproterozoic basin in the Prydz Belt in East Antarctica and its implications for Gondwana assembly and ultrahigh temperature metamorphism. Precambrian Research, 161, 355–388. Meert, J. G., 2003. A synopsis of events related to the assembly of eastern Gondwana. Tectonphysics, 362, 1–40. Meert, J. G., and Lieberman, B. S., 2008. The Neoproterozoic assembly of Gondwana and its relationship to the Ediacaran– Cambrian radiation. Gondwana Research, in press. Meert, J. G., and Torsvik, T. H., 2003. The making and unmaking of a supercontinent: Rodinia revisited. Tectonophysics, 375, 261–288. Meert, J. G., Van der Voo, R., and Ayub, S., 1995. Paleomagnetic investigation of the neoproterozoic Gagwe lavas and Mbozi Complex, Tanzania and the assembly of Gondwana. Precambrian Research, 74, 225–244. Moores, E. M., 1991. Southwest U.S.-East Antarctic (SWEAT) connection: a hypothesis. Geology, 19, 425–428. Squire, R. J., Campbell, I. H., Allen, C. M., and Wilson, C. J. L., 2006. Did the Transgondwanan supermountain trigger the explosive radiation of animals on Earth? Earth and Planetary Science Letters, 250, 116–133. Stern, R. J., 1994. Arc assembly and continental collision in the Neoproterozoic East Africa Orogen: implications for the consolidation of Gondwanaland. Annual Reviews of Earth and Planetary Science, 22, 319–351. Yoshida, M., 2007. Geochronological data evaluation: implications for the Proterozoic tectonics of East Gondwana. Gondwana Research, 12, 228–241.

Cross-references Breakup of Rodinia Critical Intervals in Earth History Ediacaran Biota Origins of the Metazoa Snowball Earth Trace Fossils: Neoproterozoic

GREAT OXYGENATION EVENT (GOE) The term “Great Oxygenation Event” (GOE, also known as “Great Oxidation Event,” “oxygen catastrophe,” or “oxygen crisis”) describes a critical environmental change in Earth history that resulted from the appearance of diatomic oxygen (O2), a waste product of oxygenic photosynthesis, in the atmosphere. The GOE occurred around 2.4 billion years (Ga) before present, close to the boundary between the Archean and the Proterozoic (2.5 Ga). For details, please refer to “Critical Intervals in Earth History.”

GREEN ALGAE Green algae are photosynthetic eukaryotes with simple plastids (derived from primary endosymbiosis) that are mostly microscopic and rarely more than a meter in greatest dimension. The approximate 6,000 extant species show an enormous diversity of growth habit and fine details of their cellular architecture. Green algae occur as unicellular or colonial, microscopic or macroscopic, motile planktonic, as well as benthic attached forms. Some (e.g., Dasycladales) are important carbonate producers in aquatic environments, others live in symbiotic associations with fungi to form lichens in terrestrial settings. Green algae living as endosymbionts inside heterotrophic organisms are a common phenomenon. The order Charales, in which full tissue differentiation occurs, is considered to include the closest relative of higher plants. For details, please refer to entry “Algae (Eukaryotic).”

GUILD Metabolically related populations, e.g., sulfate-reducing microorganisms. See entry “Microbial Communities, Structure, and Function” for further reading.

H

HABITAT The physical location or dwelling place of a particular organism, where an individual ecotype can be found or isolated from. See entry “Microbial Communities, Structure, and Function” for further reading.

HALOBACTERIA – HALOPHILES Helga Stan-Lotter University of Salzburg, Salzburg, Austria

Definition Halobacteria (plural), halobacterium (singular). Colloquial for halophilic archaebacteria (or archaea), which are single-celled life forms. Synonyms: Haloarchaea (plural), haloarchaeon (singular). Halobacterium. The name of a genus of the archaeal family Halobacteriaceae (Grant, 2001). Halophilic. Requires a salt-rich environment for growth and survival. Introduction Saline waters dominate the earth, with the oceans holding 97% of the planet’s water (a total of 1.338
109 km3; http://ga.water.usgs.gov/edu/earthwherewater.html). Saline inland seas, groundwater, and saltwater lakes hold another 0.9–0.94%, which exceeds the volume of the world’s available freshwater (1.1
107 km3). The definition of a hypersaline environment is one that possesses a salt concentration greater than that of seawater (>3.5% w/v). Water-containing environments are usually described as thalassohaline or athalassohaline (see

Chapters “Saline Lakes,” “Soda Lakes”). Thalassohaline waters are marine-derived and therefore contain, at least initially, the seawater composition; with increasing evaporation, the concentration of various salts is changing. Athalassohaline waters may also receive influx of seawater; however, the chemical composition is mainly determined by geological, geographical, and topographical parameters. Interactions between halophilic microorganisms and geological materials have been recognized early in the history of mankind and continue to impact human endeavors, including the exploration of energy sources, the storage of waste, the search for extraterrestrial life (see Chapter “Deep Biosphere of Salt Deposits”). For example, the evaporation of seawater for the purpose of making salt and the concomitant development of red color of brines, which is due to pigmented microorganisms, have been described already 2,700 years ago (Oren, 2002). It was also observed early that the presence of microorganisms in solar salterns enhances the precipitation and the yield of salt (Oren, 2002). Halite, the mineral of sodium chloride, primarily forms from the evaporation of seawater. Subsurface deposits of halite occur in many areas and are far more abundant than was previously recognized. The advent of widespread oil drilling revealed a frequent association of ancient salt deposits and the presence of the biogenic energy sources, oil and methane (Sassen et al., 1994). The potential impact of viable halophilic microorganisms in salt deposits (see Chapter “Deep Biosphere of Salt Deposits”), some of which are being used as storage sites for long-lived radioactive isotopes such as transuranic wastes, has been the focus of much research. The striking discovery of extraterrestrial halite in meteorites (see Chapter “Deep Biosphere of Salt Deposits”) and recently, in salt pools on Mars (Osterloo et al., 2008) make a search for halophilic microorganisms on other planets or moons plausible.

Joachim Reitner & Volker Thiel (eds.), Encyclopedia of Geobiology, DOI 10.1007/978-1-4020-9212-1, # Springer Science+Business Media B.V. 2011

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Saline environments and their inhabitants Organisms occur within a range of salinities, from distilled water to saturated salt solutions, and may be loosely classified as non-, slightly, moderately, or extremely halophilic, requiring about 0.2 M, 0.2–0.85 M, 0.85–2.5 M, and 2.5–5.2 M NaCl, respectively, for growth. The largest saline environment, seawater, contains 31–38 g of total dissolved salts per liter, nearly 80% of which is NaCl. Hypersaline environments are characterized by higher salinities than seawater. The oceans are inhabitated by a great diversity of organisms; when seawater is evaporating, the biological diversity decreases with increasing concentration of salt. Macroscopic organisms such as salt-tolerant fish (e.g., Cyprinodon variegatus and Menidia beryllina) are no longer found at concentrations of more than about 11% salt (1.88 M NaCl); copepods (fleas), turbellarian worms, and some rotifers tolerate 15–17% salt. Interestingly, several invertebrates such as brine shrimp (Artemia salina) or brine flies (Ephydra cinerea) are capable of survival at NaCl concentrations of 30–33% (5 M; Javor, 1989). However, the dominating organisms in saturated brines are extremely halophilic microorganisms which grow optimally in the presence of 2.5–5.2 M NaCl and are generally unable to grow in less than 1.7 M NaCl. They can occur at such high cell densities (up to 108 colony forming units per milliliter) that they cause brines to turn bright red or purplish. Figure 1 shows a commercial salt-producing facility where seawater is evaporating in a series of ponds. The colors of the pools are caused by carotenoid pigments, which are contained in the membranes of haloarchaea, or by the chlorophylls of halophilic algae or cyanobacteria. Typical haloarchaea in such communities are species of the genera Halobacterium, Halorubrum, and Halococcus, halophilic bacteria such as Ectothiorhodospira halochloris, Salinibacter ruber, cyanobacteria (e.g., Aphanothece halophytica, Phormidium sp. and Schizothrix arenaria), and green algae (e.g., Dunaliella salina and Asteromonas

gracilis). Figure 2 shows two characteristic haloarchaeal morphologies – Halobacterium salinarum NRC-1 grows as single rod-shaped cells; Halococcus salifodinae cells are roundish and tend to aggregate into large clusters. Crude solar salt contains large numbers of viable bacteria and archaea (about 105–106 colony forming units per milliliter; Oren, 2002). When using this salt for the preservation of fish, meat, and hides, damage in the form of red discoloration of the goods was noted under conditions of moisture and elevated temperatures. These observations started the early research on halophilic microorganisms in the 1920s–1930s. Only much later – in the 1980s – was it recognized that most halophiles from such sources belonged to the archaea. The haloarchaea are not pathogenic; in fact, some are used in traditionally fermented foods in Asia for human consumption (Oren, 2002). There exist other hypersaline environments which are not directly derived from seawater and are called athalassohaline (see above and Chapter “Soda Lakes”); often they are remnants of prehistoric inland lakes, such as the Dead Sea and the Great Salt Lake. Additional larger lakes include Mono Lake in California, soda lakes in Egypt and China, Lake Magadi in Kenya, and Lake Eyre in Australia. The chemical composition varies depending on the geology of the surroundings, but overall salinity is usually between 200 and 340 g/l of dissolved salts. Soda lakes are an example of naturally occurring alkaline environments which may also be hypersaline, and are characterized by the presence of large amounts of sodium carbonate and correspondingly minute concentrations of Ca2+ and Mg2+. The pH of alkaline lakes is usually in the range of 10–11; in some evaporation ponds, it can rise up to 12 (Oren, 2002). Microbial isolates from alkaline hypersaline lakes include the archaea Natronomonas sp., Natronococcus sp., Natrialba sp., and bacteria of the genera Halomonas, Chromohalobacter, Spirulina (the latter is known as nutrient for humans and animals) and others still to be characterized (Javor, 1989; Oren, 2002).

Halobacteria – Halophiles, Figure 1 Lagoons of a commercial salt-producing facility in Namibia, close to the ocean. Red-colored lagoons contain mainly haloarchaea; green color is due to halotolerant algae or cyanobacteria.

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Halobacteria – Halophiles, Figure 2 Scanning electron micrograph of cells of Halobacterium salinarum NRC-1 (left) and Halococcus salifodinae DSM8989T (right). Bars, 0.5 mm. (Photographs were taken by C. Frethem (left) and G. Wanner (right).)

From the Deep Lake in the Vestfold Hills of Antarctica, a psychrophilic archaeon, Halorubrum lacusprofundi, was isolated. Deep Lake has a salinity that is ten times higher than that of seawater (3.6–4.8 M). The lake remains ice-free throughout the year, with water temperatures fluctuating between +10 and 15 C and remaining below 0 C for 8 months of the year. The microbial diversity in Deep Lake is extremely low and is dominated by haloarchaea, with apparently low numbers of Dunaliella sp. and bacteria (of the beta- and gamma-proteobacteria) present in the sediment (Cavicchioli, 2006). Less explored and unusual hypersaline habitats are represented by anoxic brines near the bottom of the sea, for example in the Mediterranean deeps, which contain saline brines almost saturated with magnesium chloride, and where a wide diversity of novel archaea and bacteria was detected (van der Wielen et al., 2005). Another environment for halophilic microorganisms are ancient salt deposits; recent reports have described the presence of haloarchaea in Triassic and Permian salts (summarized in McGenity et al., 2000; Stan-Lotter et al., 2004). These viable microorganisms were consistently isolated from surface-sterilized crystals obtained from freshly blasted rock salt or deep drilling bore cores, suggesting that these were survivors from the original depositional event (see Chapter “Deep Biosphere of Salt Deposits”). Finally, halophilic viruses have been detected, which were released from bacteria and archaea; in addition, free virus particles were found by electron microscopical examination of the Dead Sea and in Spanish saltern ponds (Oren, 2002). The viruses (also called bacteriophages) were equally dependent as their hosts on high concentrations of salt for their structural stability. There is little information on the role of viruses in natural environments; suggestions which were made involve ecological functions in controlling the sizes of microbial communities (Oren, 2002). A website with pictures of haloviruses from

the laboratory of M. Dyall-Smith is available: http://www. microbiol.unimelb.edu.au/people/dyallsmith/research/ haloviruses/

Classification and genomics Halophilic means salt-loving and this term, often as the prefix hal- or halo-, has been in use since the 1930s. Therefore, members of the bacteria as well as the separate prokaryotic group archaea, which was identified only in the 1970s, may contain such prefixes in their names. Some confusion may arise from this nomenclatural convention, since, for example, the genus with the name Halobacterium belongs to the halophilic archaea (or haloarchaea), not to the bacteria. The number of recognized haloarchaeal genera has increased to 26, according to the International Committee on Systematics of Prokaryotes (http://www.the-icsp.org) and the number of validated species at this time is 86. Main taxonomic criteria for the identification and recognition of haloarchaea are the sequence of the 16S rRNA genes and the composition of their characteristic membrane polar lipids, which are unique ether-linked phosphoglycerides (see Fendrihan et al., 2006, for a recent overview). The complete list of required and recommended criteria for the formal identification and recognition of haloarchaeal species was proposed by Oren et al. (1997). For halophilic bacteria belonging to the large family Halomonadaceae, a similar list of specific characteristics, which has been revised recently, can be consulted (Arahal et al., 2007). Five genomes of haloarchaea have been completely sequenced, that of Halobacterium salinarum NRC-1, the related strain Halobacterium salinarum R1, Haloarcula marismortui, Natronomonas pharaonis, and Haloquadratum walsbyi, and one genome of a halophilic bacterium (Salinibacter ruber). The website http://www. ncbi.nlm.nih.gov/sites/entrez lists all genome projects, whether complete or still in progress, including references

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and detailed descriptions. Comparisons of genomes continue to provide insights about similarity and differences between archaea and bacteria and about the early evolution of prokaryotes. They have, for example, proven that the adaptation mechanism of keeping a high internal salt concentration (see next paragraph) is not unique to archaea, as was thought before, but is used also by Salinibacter ruber; this common property as well as some other characteristics suggests the possibility of early lateral gene transfers between prokaryotes (Velieux et al., 2007).

Adaptations to high salinity Most halophiles are unable to survive outside their highsalt native environments. Some species of the haloarchaea are so fragile that when placed in distilled water they lyse within minutes from the change in osmotic conditions (Grant, 2001). This reflects a general adaptation of the cells’ entire intracellular machinery to high concentrations of cations. Halophilic and halotolerant organisms have to expend energy to exclude salt from their cytoplasm to avoid protein aggregation (“salting out”). Two different strategies are known to be used to prevent desiccation through osmotic movement of water out of their cytoplasm, and both work by increasing the internal osmolarity of the cell. In the first, organic compounds, termed “compatible solutes,” are accumulated in the cytoplasm; this strategy has been found in many halophilic bacteria and also in some archaea, yeasts, algae, and fungi (Galinski, 1995). Compatible solutes are neutral or zwitterionic and include amino acids, sugars, polyols, betaines, and ectoines, as well as derivatives of some of these compounds; they can be synthesized de novo or taken up from the environment (Galinski, 1995; Grant, 2004). The second adaptation involves the selective import of K+ ions into the cytoplasm, which can reach internal concentrations of 5 M. This adaptation is restricted to the moderately halophilic bacterial order Halanaerobiales, the haloarchaeal family Halobacteriaceae, and the extremely halophilic bacterium Salinibacter ruber (Oren, 2002). Most proteins of haloarchaea and also of Salinibacter contain a large excess of acidic amino acids (glutamate and aspartate) and a low content of basic amino acids (lysine and arginine; Lanyi, 1974; Oren, 2002); these features are thought to represent a specific adaptation to high levels of salt (Mevarech et al., 2000; Oren, 2002). In contrast, the action of compatible solutes is much less specific, with little or no adjustment of intracellular macromolecules required, which is in line with the observation that these compounds act as general stress protectants against heating, freezing, and drying (see Oren, 2002, and references therein). Conclusions Halophilic organisms occur in all major groups on the tree of life (see Oren, 2007) and even halophilic viral representatives are known. Halophilic microorganisms in particular

may be found on earth in almost every hypersaline environment – tropical to polar, terrestrial to submarine, acidic to alkaline, or aerobic to anaerobic (Javor, 1989). Initially a little-studied group of microorganisms, halophilic bacteria and archaea have proven to be of interest for life sciences and earth sciences as well; their investigation has resulted in major insights for molecular biology, genomics, biogeochemistry, and – most recently – astrobiology.

Bibliography Arahal, D. R., Vreeland, R. H., Litchfield, C. D., Mormile, M. R., Tindall, B. J., Oren, A., Bejar, V., Quesada, E., and Ventosa, A., 2007. Recommended minimal standards for describing new taxa of the family Halomonadaceae. International Journal of Systematic and Evolutionary Microbiology, 57, 2436–2446. Cavicchioli, R., 2006. Cold-adapted archaea. Nature Reviews. Microbiology, 4, 331–343. Fendrihan, S., Legat, A., Gruber, C., Pfaffenhuemer, M., Weidler, G., Gerbl, F., and Stan-Lotter, H., 2006. Extremely halophilic archaea and the issue of long term microbial survival. Reviews in Environmental Science and Bio/technology, 5, 1569–1605. Galinski, E., 1995. Osmoadaptation in bacteria. Advances in Microbial Physiology, 37, 272–328. Grant, W. D., 2001. Genus I. Halobacterium Elazari-Volcani 1957, 207, AL emend. Larsen and Grant 1989, 2222. In Boone, D. R., Castenholz, R. W., and Garrity, G. M. (eds.), Bergey’s Manual of Systematic Bacteriology, 2nd edn. New York: Springer, Vol. I, pp. 301–305. Grant, W. D., 2004. Life at low water activity. Philosophical Transactions of the Royal Society of London. Series B, Biological Sciences, 359, 1249–1266. Javor, B. J., 1989. Hypersaline Environments: Microbiology and Biogeochemistry. Berlin: Springer. Lanyi, J. K., 1974. Salt-dependent properties of proteins from extremely halophilic bacteria. Bacteriological Reviews, 38, 272–290. McGenity, T. J., Gemmell, R. T., Grant, W. D., and Stan-Lotter, H., 2000. Origins of halophilic microorganisms in ancient salt deposits (MiniReview). Environmental Microbiology, 2, 243–250. Mevarech, M., Frolow, F., and Gross, I. M., 2000. Halophilic enzymes: proteins with a grain of salt. Biophysical Chemistry, 86, 155–164. Oren, A., 2002. Halophilic Microorganisms and Their Environments. Dordrecht, The Netherlands: Kluwer. Oren, A., 2007. Biodiversity in highly saline environments. In Gerday, C., and Glansdorff, N. (eds.), Physiology and Biochemistry of Extremophiles. Washington: ASM, pp. 223–231. Oren, A., Ventosa, A., and Grant, W. D., 1997. Proposed minimal standards for description of new taxa in the order Halobacteriales. International Journal of Systematic Bacteriology, 47, 233–236. Osterloo, M. M., Hamilton, V. E., Bandfield, J. L., Glotch, T. D., Baldridge, A. M., Christensen, P. R., Tornabene, L. L., and Anderson, F. S., 2008. Chloride-bearing materials in the southern highlands of Mars. Science, 319, 1651–1654. Sassen, R., Cole, G. A., Drozd, R., and Roberts, H. H., 1994. Oligozene to holocene hydrocarbon migration and salt dome carbonates, Northern Gulf of Mexico. Marine and Petroleum Geology, 11, 55–65. Stan-Lotter, H., Radax, C., McGenity, T. J., Legat, A., Pfaffenhuemer, M., Wieland, H., Gruber, C., and Denner, E. B. M., 2004. From intraterrestrials to extraterrestrials – viable haloarchaea in ancient salt deposits. In Ventosa, A. (ed.), Halophilic Microorganisms. Berlin: Springer, pp. 89–102.

HISTOLOGY

van der Wielen, P. W., Bolhuis, H., Borin, S., Daffonchio, D., Corselli, C., Giuliano, L., D’Auria, G., de Lange, G. J., Huebner, A., Varnavas, S. P., Thomson, J., Tamburini, C., Marty, D., McGenity, T. J., and Timmis, K. N., and BioDeep Scientific Party, 2005. The enigma of prokaryotic life in deep hypersaline anoxic basins. Science, 307, 121–123. Velieux, F., Madern, D., Zaccai, G., and Ebel, C., 2007. Molecular adaptation to high salt. In Gerday, C., and Glansdorff, N. (eds.), Physiology and Biochemistry of Extremophiles. Washington: ASM, pp. 240–253.

Cross-references Archaea Astrobiology Deep Biosphere of Salt Deposits Extreme Environments Hypersaline Environments, Terrestrial Saline Lakes Soda Lakes

HAPTOPHYTES Haptophytes (Haptophyta, Prymnesiophyta) are unicellular chlorophyll a and c containing algae with complex plastids derived from secondary endosymbiosis (“red lineage”) (see entry “Symbiosis”). They occur principally as solitary free-living motile cells that possess two smooth flagella, unequal in length in the Pavlovophyceae and more or less equal in the Prymnesiophyceae. Haptophytes inhabit littoral, coastal, and oceanic waters and are important primary producers in many aquatic environments. Within the haptophytes, the so-called coccolithophores (coccolithophorids) evolved the ability to control the intracellular calcification onto organic plates and the assembly of the mature calcium carbonate (calcite) scales (coccoliths) at the cell surface. These forms have been significantly contributing to the deposition of calcium carbonate in marine waters since the Mesozoic, today accounting for about a third of the total marine CaCO3 production. In modern and ancient sediments, haptophyte-derived organic matter contributions can be specified by distinctive lipid biomarkers (alkenones), whose unsaturation patterns also provide a paleo-thermometer for past surface water temperatures (see entry “Biomarkers (Molecular Fossils)”). For further details, please see entry “Algae (Eukaryotic).”

HEAVY METALS Please refer to entries “Acid Rock Drainage,” “Arsenic,” “Biomining (Mineral Bioleaching, Mineral Biooxidation),” “Copper,” “Geomycology” “Metalloenzymes,” “Metals, Acquisition by Marine Bacteria,” “Ores, Microbial Precipitation and Oxidation,” “Zinc.”

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HISTOLOGY Michael Gudo1, Gerta Fleissner2, Guenther Fleissner2 1 Morphisto Evolutionsforschung und Anwendung GmbH, Institut für Evolutionswissenschaften, Frankfurt am Main, Germany 2 Goethe-University Frankfurt, Frankfurt a. M., Germany

Definition Histology is the biological science concerned with the minute or microscopic structures of cells, tissues, and organs of animals, plants, and fungi, in relation to their function, and any organismic constructions produced by living individuals or symbiotic assemblies of living organisms. Biopsy is the sample of an organism or part of a tissue that is processed for histological investigations. Fixation serves the inhibition of all vital processes by special chemicals that also preserve and prepare organic matter for further histological or morphological investigations. Mineralization is the process by which organic components of organisms are impregnated or replaced by inorganic material and converted to mineral matter. Staining is the chemical reaction of particular laboratory chemicals (stains and dyes) with tissue components and materials of the samples. Introduction The term histology was coined by C. Mayer in 1819 as a part of the scientific discipline comparative anatomy (Mayer, 1819). Histological investigations, concerned with the minute structures of cells, cell organelles, intraand extracellular fiber arrangements, and tissue components and organs in relation to their function, are usually done by light microscopy. Since the invention of the microscope by Leeuwenhoek in 1668 (Palm and Snelders, 1983), the early histological technology (Gerlach, 1998) has experienced major improvement by the refinement of microscopes (Bozzola and Russell, 1999) and new methods of tissue preservation, sectioning devices, stains, and staining methods. Histochemical methods now permit to identify and localize intracellular components and enzymes within distinct and isolated samples (e.g., Bancroft and Gamble, 2007; Böck, 1989; Horobin and Kiernan, 2002; Mulisch and Welsch, 2010). Therefore, the studied structures, tissues, complete organisms, or any organismic compound have to be preserved from enzymatic decay, and then cut into sections of just a few microns thickness. The histological sections can vary between ultra thin sections (40–80 nm), semi thin sections (0.1–3.0 µm), thin sections (3–20 µm), and thick sections (more than 20 µm). While ultra thin sections could be investigated only by transmission or scanning electron microscopy (TEM), the others can be investigated with any light microscope. Afterward, these sections are

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processed by various histochemical and biophysical techniques according to the aim of the study. Geobiological samples usually do not consist of organic soft material, alone, like the above mentioned biological materials, but they are often composites of hard inorganic and mineralized organic material, with nearly all biological components converted to inorganic matter. Some geobiological samples are compositions of both, organic and inorganic materials, such as bio films growing on hard substrates, or sessile organisms that adhere to hard substrates like corals, sponges, lichens, or fungi. These samples therefore can seldom be histologically processed like biological material via paraffin embedding and sectioning with traditional microtomes. Special procedures and mechanical devices (like saw microtomes, annular saws or cutting/grindings systems) are necessary to produce sections of geobiological samples (e.g., Hoffmann et al., 2003). By means of special polishing machines, like the ultramill or lapping machines, sections or slices of these sections can be studied by distinct microscopical methods, for example under fluorescent light, X-ray-mappings and topographic element analyses (Janssens et al., 2000) can identify the outline of the biological compounds and their environment, and they may also serve as a biomarker; harsh biochemical treatment can isolate the organic from the inorganic components, but often destroy the original context. Histological analyses may provide tools to identify the biological components even if their former structural context has decayed (Wrede et al., 2008), or help to analyze particular processes or compositions (Neuweiler et al., 2007). Principally these results must be “calibrated” by means of histological investigations of recent tissues in order to understand organ structure and its function (Young and Heath, 2000). This entry reviews the state of the histological methods and offers the modifications necessary to topographically investigate geobiological samples. For clarity, this latter information is given in italics.

Methods Different from biochemical investigations in the broadest sense, histological analyses are concerned with samples, where the structures and their topographical context are preserved or known before biopsies are taken and only in a controlled way modified by procedures according to the specific aim of the study. For histological processing of tissues or any other organic compounds, all vital functions have to be stopped by fixation, in order to preserve a distinct structure within or more often isolated from its natural environment. In geobiology this process often is “history,” but must be known to understand the given results of further analyses. Usually, the biological samples are embedded into a special medium which provides particular mechanical properties that allow sectioning in the appropriate thickness. The sections are then placed on glass slides or cover

slips, afterwards stained by special chemicals and dyes that have affinities to particular tissues or histological structures. The staining allows the differentiation of cell structures, various types of cell surface and skeleton assemblies, filaments or fibrillar components (e.g., collagenous fibers, elastic fibers, or reticular fibers), tissues, and even complete organs or pathological and genetic modifications of the various structures. Under the microscope the results can be observed and – most often after a thorough comparison to reference samples – evaluated and interpreted. In geobiology, the selective staining of minerals prevails, although protein staining has been successful in various studies. The major problem in geobiology is that mineralized samples are too hard for conventional section procedures and therefore special preparation methods are necessary (see also below, paragraph “Embedding”). Accordingly, often only the surface of sectioned samples is accessible to the staining which limits the differentiations of components. Structural analyses of biological compounds have to refer to similar structures or organs histologically known from recent specimen. Therefore, histology in geobiology has to refer to a biological histology (Robenek, 1995). Specialized methods are available which either demineralize the sample for showing the biological contents or dissolve the organic leftovers to investigate their “casts” in the inorganic surroundings. Often the resulting histologically processed specimens can be biophysically investigated, and topographic X-ray element mappings (Janssens et al., 2000) are often the method of choice to serve as biomarkers.

Fixation and infiltration The process of fixation and infiltration of biological specimens is most crucial for the results and aims to stop any autolytic and bacterial decomposition. The most commonly used fixative is formalin; however, other fixatives (e.g., Bouin, Carnoy, Paraformaldehyde, potassium dichromate, chromic acid, or mercury-fixation) are necessary depending on the planned subsequent protocol. The choice of the fixative determines the results, since every fixative has assets and drawbacks for the preservation of particular histological structures and especially the binding sites for differential staining, e.g., in immunohistology. For biophysical analyses, the metalfree fixatives and also metal-free pre- and post-processing protocols are essential. The fixation process, which is a chemical reaction, has to be controlled and stopped after a particular time that depends on type and size of the tissue, but also on environmental factors like temperature and air pressure. Afterwards the fixative must be removed following a special protocol, which takes care not to destroy structural characteristics (Böck, 1989). Prior to embedding, the sample has to be slowly transferred to the final substance via several intermediate steps mixable with the respective embedding medium, e.g.,

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for the paraffin/wax embedding dehydrants (ethanol, isopropanol, and methanol) and clearants (xylene, toluene, and xylene-replacing clearants). In case of aerated tissues, it may be necessary to perfuse the sample with a prefix and process fixation and embedding continuously in a vacuum with fluid movement for a proper impregnation of all layers of the specimen. In geobiology, fixation and conservation also plays a crucial role, since these chemical processes have to be balanced between the necessities for the mineralized and organic components of a geobiological sample (biofilms or water containing fossils; e.g., Desrochers et al., 2007). Every fixative generates chemical reactions, which sometimes may degrade or destroy either the organic or the mineralized components. Accordingly, empirical studies have to be performed before crucial samples are processed in order to ascertain whether the fixative may cause any problems for either the soft or hard tissues of a particular type of samples.

Embedding Embedding of histological material is performed in order to provide a mechanical stable and hard entity, which can be sectioned to various coherent thin sections. The sections should enable a spatial reconstruction of the original sample after differentiating analyses. The composition of the specimen and the histological program are decisive for the embedding methods of choice. The most common material for tissue embedding is paraffin (waxembedding), also embedding in celloidin and resin. If the tissues under study are more or less soft (which means without any calcareous or other mineralized structures), the paraffin embedding is the best choice. Celloidin embedding takes much time and is useful for very sophisticated questions (e.g., embedding of complete brains or very large organs). Plastic embeddings are the best choice if the biopsies contain mineralized structures, such as calcium carbonate, calcium phosphate, and silicate, or if organismic components adhere to rocks or other hard substrates, like for example biofilms. Therefore, these latter embedding methods lend themselves for geobiological samples, which then can be processed like biological material. For resin embedding (Howat et al., 2007) one can choose between epoxy-embedding, methyl-methacrylate-embedding, or glycol-methacrylate-embedding. Various products are commercially available (Obenauf et al., 2007), all of them have well-documented assets and drawbacks. The most drawbacks of plastic embeddings can be seen in their high costs of time and money. On the other hand, resin embedded samples can be sectioned, sawed, milled, or polished by use of various microtomes, saws, milling, or polishing machines. Sectioning Whole mount samples can only be investigated from their outer surface. Analyses inside the samples need dissection

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with preferably minor destruction of the structural context. In general, penetration of staining chemicals is enhanced afterwards, and microscopic and biophysical inspection is improved as the spatial resolution corresponds to the thickness of the section. Several problems may arise from inappropriate section procedures:  The samples may be too brittle and break. This effect is

reduced by pre-treatment procedures like decalcification or demineralization, also choosing another type of embedding medium may help possibly combined with different section techniques, e.g., a vibrating knife or sample cooling by a special refrigerant device.  The sections are heavily distorted. A proper selection of the microtome, cutting angle, and cutting velocity may improve the result. The sections may stretch, for example, after floating on a warm water bath about 10 C below the melting point of the embedding wax.  The requested thickness requires different blades, which cannot be used “universally,” even if section series of varying thickness are necessary for the protocol. For example, thick sections need a hard metal blade, while semithin- and ultrathin sections can be produced only by means of glass or diamond knives. Biological samples, which are embedded in paraffin or resin, can generally be sectioned by a special device, a microtome. Some microtomes are equipped with a motor-driven support, both for controlling the section thickness as well as the force of the blade. Several types of microtomes are available, each optimized for variably embedded samples: Paraffin embedded samples can be sectioned by sled microtomes or rotary microtomes equipped with metal knives. Resin embedded samples need microtomes which have a special retraction mechanism, to avoid mechanical damage of the knife and the sample. The blades are either manufactured of especially hardened steel for sections between 5 and 30 µm, for semi thin sections (2–5 µm) and ultra thin sections (about 40–100 nm) glass or diamond knives must be used. Hard materials, like mineralized geobiological probes, cannot be sectioned by conventional microtomes unless the samples have been pre-treated: decalcification or demineralization by different chemicals. While these procedures may cause uncontrolled damage, when applied to the unsectioned samples, preferably control material should be preserved to evaluate the original structural context. Therefore, careful special section devices like diamond milling, saw microtomes, annular saws, or section/ grinding-systems especially optimized for this purpose should be applied. About 30 µm thick sections of rather large samples can be achieved by these methods. The sections or grinded pieces can then be further processed, and also partly preserved for subsequent evaluation. Another method, available since several years, is the laser microtome which may contact free isolate minute samples out of the otherwise untreated entity. The samples must neither be pre-treated nor embedded. Section thickness ranges between 10 and 100 µm. Special

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devices allow even for the isolation of minute tissue pieces like a single cellular nucleus. A satisfying section quality requires a long time of experience especially to decide for the most appropriate procedure. While continuously new embedding products are commercialized, only little has changed concerning the basic techniques of standard microtomy, which is taught as a component part of the education of laboratory and technical assistants.

Mounting For further histological processing, the sections must be mounted, most often on glass slides or on grids coated with a special synthetic film. The major difficulty is the choice of the best possible sticking procedures in order to prevent both, a floating resp. a loss of the sections during the staining protocol and undesired background staining by the glue. While before often unreliable “home-made” coating procedures, e.g., by chromic sulphate or polylysine, were used, now various types of pretreated glass slides are commercially available, they have been optimized for selected histological procedures. Thick sections, e.g., sections that are produced by saw microtomes or cutting/grinding-systems, have to be glued to object slides by special sticking resins that, e.g., harden with ultraviolet light; in this aspect, histological sample preparation is quite similar to the preparation of geological thin ground-sections. Biophysical analyses of geobiolocal samples requires mounting according to the planned technique (see list of Handbooks of the suppliers). In, for example, X-ray investigations (micro fluorescence analyses), the mounting must be free of auto fluorescence; the sometimes long lasting measuring intervals need a stable non-vibrating mechanical stability. Staining Multiple histological staining methods allow identifying cellular structures, tissue components, and organ systems, also enzymatic processes and involved macromolecules. It is even possible to clearly differentiate between organic and inorganic material as well as decaying from living material by selective marking of minerals and molecules. Stainings can be performed by use of natural stains and dyes or by chemicals often designed for certain projects. New fields of histological analyses have been launched by immunohistochemical stainings combined with fluorescent markers which are very selectively binding to particular proteins. Differential and double stainings, also section series with alternatively stained sections, have revealed important insight into basic cellular processes, information processing, and also pathological resp. genetic changes. The standard protocols are well documented including their modifications for the application to tissues of various origins.

For any staining it is necessary that the structures that should be stained are free of any embedding media. This means that for standard-histology the paraffin has to be removed by special procedures so that the staining chemicals can react with the tissues. If samples are embedded in resins, this process is more crucial and difficult; several resins cannot be removed and accordingly only the surface of the section provides reactive structures for staining. The staining of cellular structures and tissues is based on chemical affinities of the stains with particular tissues resp. tissue components. In many cases, a particular pretreatment is necessary for adequate results. Pretreatments may be, for example, mordants, which modify the stain so that it reacts with the tissue or modifies the tissues so that the stain can react with the tissue. In some cases mordants generate a reaction that transformed the water soluble stain into a water- and alcohol-stable lacquer. This chemical reaction can, for example, be induced by metal ions like tungsten or molybdenum or by alkali ions like lithium, sodium, or barium (Böck, 1989; Burck, 1988). Stainings are usually composed of three steps: in the first step cellular structures like nuclei, etc., are stained. In the second step cell plasma and some extracellular structures are stained, and in the third step the surrounding matrix is stained. As a result of the staining, the nuclei, cells, various fibers, and other structures or chemical components in a tissue have particular colors that allow differentiation and several diagnostics. Some stainings also provide particular optical characteristics to tissues; they may for example provide polarizing effects to the tissue. Other procedures provide fluorescence-effects or particular ultra-violet-effects of the polished surface and allow further investigations (Figures 1 and 2). While traditional stainings induce a chemical reaction within tissues and may stain in an unselective way (Horobin and Kiernan, 2002), immunohistology (Polak and Noorden, 2003) selectively marks specific epitopes, proteins, and molecules bound to membrane or subcellular structures. Traditional and immunohistochemical analyses under the light microscope meet the limits of resolution at magnifications of about 1,200
; therefore, immunohistological methods have been upgraded for analyses under the TEM, either by pre-embedding or post-embedding staining protocols. While traditional stainings are used for topographic and micro anatomical investigations, immunhistological stainings are used to understand the functional context of molecular processes and to allocate the regions in which these processes take place in an organism or in a tissue. These refined immunohistological staining methods have also been applied to sections of geobiological samples. The analyses could demonstrate biological compounds, thus serving as biomarkers, and in some cases even disclose the genetic origin of the fossilized organisms.

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Histology, Figure 1 Cross section of a human long bone under different microscopic optics (thin-ground section of a nondemineralized sample). (a) Under through light with minimal color saturation; (b) same illumination with maximum color saturation; (c) differential interference contrast (After Nomarski). Already without staining procedures characteristic structural details and varying densities can be identified. (a–c same magnification.)

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Histology, Figure 2 Section series through a chrondral ossification zone of a demineralized porcine long bone after various differential histological staining procedures (Protocols according to Bo¨ck et al., 1989; Young and Heath, 2000). (a) Masson Goldner A; (b) Masson Goldner L; (c) Masson Goldner ResF A; (d) Masson Goldner ResF L; (e) Crossmon A; (f) Crossmon L; (g) Azan after Geidis; (h) Movat Pentachrome; (i) Safranin Fastgreen. The layers of young bone (upper area), cartilage (middle area), and secondary ossification area (below) can be analyzed concerning their different structural details (a–i same magnification).

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Conclusion Geobiology is defined as field of science which analyses the interaction of the environment with living matter. Mainly fossilized samples are investigated concerning their systematic relationship and – even more – the origin of this specific form of life. Research results can find their application not only in a better understanding of our life on earth, but also in exploring putative extraterrestrial biology. Another aspect is the transfer of basic sciences to various aspects of biomedical research like the synthesis or degradation of bones and teeth or the control of biogenic mineralization processes. All these research topics cannot be planned and performed or adequately evaluated without a thorough knowledge of the biological structures mainly based on the comparison of living and fossilized specimen. Histology delivers the methods to topographically analyze organisms, organs, tissues, and even molecules. Their modifications by environmental or biogenic processes can be recognized by the investigation of their complex interaction. Perspectives Histological investigations provide a huge field of research for biology and geobiology. Both fields have to be combined, since geobiological samples often can only be interpreted with reference to known biological samples. Some examples are histological sections of dinosaur bones or teeth that have to be compared to bones and teeth of recent organisms, others are the discoveries at special fossil-sites, like the Messel pit, the Burgess-Shale, the rocky Hunsrück region, etc., which may provide interesting information about fossilized microscopic structures. And even the investigation of recent samples like biofilms or coatings is of major relevance to understand their composition and emergence in order to discriminate between biogenic and abiotic origin. Summary Histology provides methods to topographically investigate the structure of cells, tissues, and organs down to the molecular level, especially in geobiology as reference to recognize, for example, the phylogenetic origin and age and also as a basis to analyze biological in comparison to bio geological turnover processes, which induce structural changes via environmental and climatic conditions. Bibliography Bancroft, J. D., and Gamble, M. (eds.), 2007. Theory & Practice of Histological Techniques, 6th edn. Edinburgh: Churchill Livingstone. Böck, P. (ed.), 1989. Romeis Mikroskopische Technik. München/ Wien/Baltimore: Urban & Schwarzenberg. Bozzola, J. J., and Russell, L. D., 1999. Electron Microscopy, 2nd edn. Boston: Johns and Bartlett.

Burck, H. C., 1988. Histologische Technik, 6th edn. Stuttgart: Thieme. Desrochers, A., Bourque, P.-A., and Neuweiler, F., 2007. Diagenetic versus biotic accretionary mechanisms of bryozoan-sponge buildups (Lower Silurian, Anticosti Island, Canada). Journal of Sedimentary Research, 77, 564–571. Gerlach, D. (ed.), 1998. Die Anfänge der histologischen Färbung und der Mikrophotographie – Josef von Gerlach als Wegbegleiter. Thun und Frankfurt am Main: Verlag Harri Deutsch. Hoffmann, F., Janussen, D., Dröse, W., Arp, G., and Reitner, J., 2003. Histological investigation of organisms with hard skeletons: a case study of siliceous sponges. Biotechnic and Histochemistry, 78(3–4), 191–199. Horobin, R. W., and Kiernan, J. A., 2002. Conn’s Biological Stains, 10th edn. Oxford: Bios Scientific. Howat, W. J., Warford, A., Mitchell, J. N., Clarke, K. F., Conquer, J. S., and McCafferty, J., 2007. Resin tissue microarrays: a universal format for immunohistochemistry. Journal of Histochemistry and Cytochemistry, 55, 21–24. Janssens, K. H. A., Adams, F. C. V., and Rindby, A. (eds.), 2000. Microscopic X-ray Fluorescence Analysis. Chichester: Wiley. Mayer, C., 1819. Ueber Histologie. Bonn. Cited in: Gräfe, C. F., Hufeland, C. W., Link, H. F., Rudolphi, K. A., Siebold, E. v. (eds.), 1828. Encyclopädisches Wörterbuch der medicinischen Wissenschaften, Band 2. Berlin: J.W. Boike. Mulisch, M., and Welsch, U. (eds.), 2010. Romeis Mikroskopische Technik. Heidelberg: Spektrum Akademischer Verlag. Neuweiler, F., Daoust, I., Bourque, P. A., and Burdige, D. J., 2007. Degradative calcification of a modern siliceous sponge from the Great Bahama Bank, The Bahamas: a guide for interpretation of ancient sponge-bearing limestones. Journal of Sedimentary Research, 77, 552–563. Obenauf, R. H., Martin, J., Bostwick, R., et al. (eds.), 2007. Handbook of Sample Preparation and Handling, 10th edn. Metuchen: SPEX Certiprep. Palm, L. C., and Snelders, H. A. M. (eds.), 1983. Antoni van Leeuwenhoek 1632–1723. Studies on the life and work of the Delft scientist commemorating the 350th anniversary of his birthday. Amsterdam: Rodopi. Polak, J. M., and Van Noorden, S., 2003. Introduction to Immunocytochemistry, 3rd edn. Oxford: Bios Scientific. Robenek, H. (ed.), 1995. Mikroskopie in Forschung und Praxis. Darmstadt: Git Verlag. Ruzin, S. E., 1999. Plant Microtechnique and Microscopy. Oxford: Oxford University Press. Schüler, D. (ed.), 2006. Magnetoreception and Magnetosomes in Bacteria. Berlin: Springer. Wrede, C., Heller, C., Reitner, J., and Hoppert, M., 2008. Correlative light/electron microscopy fort the investigation of microbial mats from Black Sea Cold seeps. Journal of Microbiological Methods, 73, 85–91. Young, B., and Heath, J. W., 2000. Wheather´s Functional Histology – A Text and Colour Atlas. Edinburgh: Churchill Livingstone.

Cross-references Animal Skeletons, Advent Animal Biocalcification, Evolution Biofilms Biomarkers (Molecular Fossils) Calcification Calcified Cyanobacteria

HOT SPRINGS AND GEYSERS

Carbonate Environments Fluorescence-In-Situ-Hybridization (FISH) Immunolocalization Metalloenzymes Metals, Acquisition by Marine Bacteria Microbial Biomineralization Microbial Communities, Structure, and Function Microbial Mats Organomineralization Scanning Probe Microscopy (Includes Atomic Force Microscopy) Sponges (Porifera) and Sponge Microbes

HOT SPRINGS AND GEYSERS Brian Jones1, Robin W. Renaut2 1 University of Alberta, Edmonton, Alberta, Canada 2 University of Saskatchewan, Saskatoon, SK, Canada

Definition A hot spring is a discharge of hot (>35–40 C) water from a vent at the Earth’s surface. A geyser is a hot spring characterized by intermittent, turbulent discharges of boiling water and steam. A sublacustrine hot spring is a hot spring that discharges from the floor of a lake.

Introduction A hot spring is characterized by discharge of hot water from a vent. There is, however, no universally accepted definition of “hot” and the temperature for distinguishing a “warm spring” from a “hot spring” remains contentious (Pentecost et al., 2003). In general usage, a hot spring is one with vent water temperature between about 40 C and boiling point (Renaut and Jones, 2000). It must be remembered, however, that boiling temperature changes with altitude; thus, boiling in Yellowstone National Park occurs at 92 C, whereas in New Zealand geothermal areas, which lie closer to sea level, it is at 100 C. The term “geyser” is derived from “Geysir,” located in southwest Iceland. First mentioned in historical records from 1294, the Icelandic word “geysir” means “gusher” or “one who rages.” Accordingly, a geyser is regarded as hot spring that is characterized by intermittent, turbulent discharges of boiling water and steam (Bryan, 2005, 2008). Geysers and hot springs are end members of a continuous spectrum that includes “intermittent springs” that experience periodic overflows but never erupt, and “pulsating springs,” “perpetual spouters,” or “ebullient springs” that are characterized by continual eruptions. Some hot springs discharge on the floors of lakes and in caves.

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Distribution Hot springs and geysers are found on every continent except mainland Antarctica (Waring, 1965). Many hot springs and geysers, known from historical records, have ceased activity because of natural and anthropogenically induced changes to their plumbing systems. Yellowstone National Park, USA., El Tatio located high in the Andes of northern Chile, the Taupo Volcanic Zone on the North Island of New Zealand, Kronotski National Park in Kamchatka in eastern Russia, and Iceland are the largest known geothermal fields with numerous hot springs and geysers. Bryan (2008), for example, estimated that more than 50% (500) of known geysers are located in Yellowstone National Park, with another 385 in the other four major geothermal areas. Low-enthalpy hot springs that discharge at maximum temperatures of about 35–60 C are much more common. The plumbing system The hot water discharged by a hot spring or geyser is the final, surface expression of a complex system that involves water, a heat source, and an intricate underground plumbing system. The water is derived from meteoric waters that have seeped deep into the Earth’s crust, supplemented in some regions by connate, magmatic, or metamorphic waters. Irrespective of its origin, some spring or geyser water may come from depths of 4 km where it has been magmatically heated to temperatures of >225 C. In other settings, however, meteoric waters may only penetrate to shallow depths before being expelled from a low-enthalpy spring vent. The plumbing system of a hot spring is unrestricted and allows continuous movement of water to the surface. In contrast, a geyser has a restriction within its plumbing system, commonly a sinter caprock that precludes continuous vertical flow. The configurations of these plumbing systems remain unknown because of the difficulties in accessing them. Nevertheless, schematic depictions of vast underground, water-filled chambers are probably incorrect. Instead, it seems that the plumbing systems are formed of complex networks of faults, fractures, small cavities, and strata with variable permeability. Geysers are less common than hot springs because of the special plumbing system requirements. Water trapped below a restriction in a geyser plumbing system will be heated by steam that is moving upwards from deeper parts of the system. The reservoir will then empty as the trapped superheated water is violently discharged. Such an eruption will continue until the reservoir is emptied. The magnitude, frequency, and duration of the eruption depend on reservoir size, the rate of heat transfer, and the rate at which water refills the system once emptied by the eruption. By necessity, therefore, geysers can only exist if the bedrock can withstand the pressures created by the steam and if the bedrock is permeable enough to allow rapid reservoir replenishment.

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Water acidity and composition Most springs and geysers discharge alkaline water with a pH 7. Most high-enthalpy geothermal areas (e.g., Taupo Volcanic Zone of New Zealand), however, include springs that discharge acidic waters with a pH 7 and commonly aragonite) and (or) Mg:Ca ratio (calcite < 1:1 > aragonite). Nevertheless, both calcite and aragonite are found around some hot springs and geyser vents where water temperatures are >>45 C. Spring waters discharged with high PCO2 may, through rapid CO2 degassing, quickly attain very high levels of supersaturation with respect to CaCO3. This commonly seems to be responsible for calcite and aragonite precipitation even if the waters are >75 C and have a Mg:Ca ratio of 75–80 C, include Bacteria and Archaea (Sheehan et al., 2005). Most Archaea are in the form of cocci, rods, or discs (Stetter, 1996, his Table 3). These nondistinctive morphologies coupled with the problems of laboratory culturing due to their poorly known growth requirements and nutritional needs have made species identification difficult. 16S rRNA sequencing, however, has shown that a diverse array of hyperthermophilic Bacteria, and Archaea thrive at high temperatures and extremes of pH, redox state, and salinity (Stetter, 1996; Kvist et al., 2005). Among the Bacteria, Aquifex pyrophilus and Thermotoga maritima thrive up to 95 C and 90 C, respectively. Among the Archaea, members of Pyrobaculum, Pyrodictium, Pyrococcus, and Methanopyrus survive in temperatures up to 100 C. The thermoacidophile Sulfolobus, for example, thrives in sulfur-rich hot springs at a pH of 2–3 and temperatures up to 90 C. An easy way of collecting microbes that live in the dangerous environs of hot springs/geysers is to place sterile glass slides in the water for 24–72 h. Microbes quickly colonize such substrates and remain attached once the slides are extracted from the hydrothermal waters. Microbe preservation Mineralized microbes are commonly found in precipitates that form in hot spring and geyser systems (Figure 2).

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Hot Springs and Geysers, Figure 1 Hot springs and geysers in the Taupo Volcanic Zone on the North Island of New Zealand (a, b, e, f) and El Tatio, northern Chile (c, d). (a) Prince of Wales Feathers and Pohutu during eruptive phase. Note silica precipitates on surrounding Geyser Flats. Whakarewarewa, Rotorua. (b) Margin of Champagne Pool showing exposed siliceous sinters (right) and submerged shelf covered with orange, gold- and silver-rich siliceous sinter. Water has a constant temperature of 75 C and pH of 5.4. Waiotapu geothermal area. (c) Siliceous sinter forming around margin of hot-spring pool, El Tatio. (d) Streamers of filamentous microbes in outflow channel from hot-spring pool, El Tatio. (e) General view of spring in northern part of Waiotapu geothermal area, water temperature of 80 C and pH of 2.1–2.4. Submerged reddish-brown precipitates formed largely of noncrystalline As-rich hydrous ferric oxide, poorly crystalline lepidocrocite, and crystalline jarosite. (f) Black coating around small hot spring (T = 100 C, pH = 2.0) in Kerosene Creek area, north part of Waiotapu geothermal area formed of bitumen generated by the spring system.

Such mineralization, which may involve replacement of the organic tissues and (or) encrustation around the microbes, must take place before the microbes are lost through decay. Microbe preservation is common in non-crystalline precipitates (e.g., opal-A, HFO) but rare in crystalline precipitates (e.g., calcite, aragonite, jarosite), possibly because the former generally precipitate at a faster rate than the latter and organic materials are less prone to destruction by crystal growth.

The microbes can play an active role (i.e., they mediate precipitation through modification of their microenvironment) or a passive role (i.e., they act as templates for precipitation by having nucleation sites on their surfaces) in their own mineralization. Experimental work and interpretation of naturally mineralized microbes indicate that microbes are probably passive during their own mineralization. There is, however, some evidence that some microbes may exert some influence over the initial pattern of opal-A precipitation (Jones et al., 2004).

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Hot Springs and Geysers, Figure 2 SEM photomicrographs of siliceous microbes found in Champagne Pool (a, b; T = 75 C, pH = 5.2) and Iodine Pool (c–g; T = >80 C, pH = 8.2–9.0), North Island of New Zealand. (a) Silicified filamentous microbes in flocs that circulate in waters of Champagne Pool; (b) Silicified microbes in orange siliceous sinters from shelf around Champagne Pool (see Figure 1b); (c) Silicified rod-shaped microbes; (d and e) Rod-shaped microbes encrusted with different styles of opal-A spheres; (f) Filamentous microbes encrusted with layer of very small opal-A spheres and outer layer of larger opal-A spheres; (g) Silicified filamentous microbes encrusted with opal-A spheres of various sizes.

SEM imaging commonly reveals microbes that appear to be superbly preserved (Figure 2). It is, however, very difficult to identify such microbes in terms of extant taxa because most of the taxonomically important features are lost and DNA is generally not preserved. Thus, identifications must rely solely on morphological features such as general configuration, length, width, and the presence or absence of septa. The problem is compounded by the fact that most extant taxa are defined by DNA sequencing and may not have been imaged to show their morphology.

Conclusion Hot springs and geysers are spectacular geological features that encompass a wide range of environments that are commonly characterized by extreme conditions (high T, low pH). Although opal-A, calcite/aragonite, and various Fe-rich deposits are the most common minerals precipitated from hot spring and geyser waters, these spring deposits commonly contain significant amounts of other elements, including Au, Ag, As, S, and many others. Many hot spring and geyser systems are inhabited by a diverse array of microbes (archaea, bacteria, cyanobacteria, fungi, diatoms) that flourish in the warm discharge waters. Such microbes are frequently preserved, typically in opal-A, and thereby provide some insights into the nature of the microbial communities that inhabit these complex environs. Such fossils have received

considerable attention, as they are commonly considered analogous to silicified microbes that are found in much older deposits, especially those in the Precambrian.

Bibliography Bryan, T. S., 2005. Geysers. What They Are and How They Work, 2nd edn. Missoula, MT: Mountain Press. Bryan, T. S., 2008. The Geysers of Yellowstone. 4th Edition, University Press of Colorado. Folk, R. L., 1994. Interaction between bacteria, nannobacteria, and mineral precipitation in hot springs of central Italy. Géographie physique et Quaternaire, 48, 233–246. Cady, S. L., and Farmer, J. D., 1996. Fossilization processes in siliceous thermal springs: trends in preservation along thermal gradients. In Block, G. R., and Goode, J. A. (eds), Evolution of Hydrothermal Ecosystems on Earth (and Mars?). Ciba Foundation Symposium. Chicester, UK: Wiley, pp. 150–173. Jones, B., and Renaut, R. W., 1996. Morphology and growth of aragonite crystals in hot-spring travertines at Lake Bogoria, Kenya Rift Valley. Sedimentology, 43, 323–340. Jones, B., and Renaut, R. W., 2007. Selective mineralization of microbes in Fe-rich precipitates ( jarosite, hydrous ferric oxides) from acid hot springs in the Waiotapu geothermal area, North Island, New Zealand. Sedimentary Geology, 194, 77–98. Jones, B., Renaut, R. W., and Rosen, M. R., 1997. Biogenicity of silica precipitation around geysers and hot-spring vents, North Island, New Zealand. Journal of Sedimentary Research, 67, 88–104. Jones, B., Renaut, R. W., and Rosen, M. R., 2001. Biogenicity of gold- and silver-bearing siliceous sinters forming in hot (75 C)

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anaerobic spring-waters of Champagne Pool, North Island, New Zealand. Journal of the Geological Society of London, 158, 895–911. Jones, B., Konhauser, K. O., Renaut, R. W., and Wheeler, R., 2004. Microbial silicification in Iodine Pool, Waimangu geothermal area, North Island, New Zealand: implications for recognition and identification of ancient silicified microbes. Journal of the Geological Society of London, 161, 983–993. Pentecost, A., Jones, B., and Renaut, R.W., 2003. What is a hot spring? Canadian Journal of Earth Sciences, 40, 1443–1446. Kvist, T., Mengewein, A., Manzei, S., Ahring, B. K., and Westermann, P., 2005. Diversity of thermophilic and nonthermophilic crenarchaeota at 80 C. FEMS Microbiology Ecology, 244, 61–68. Renaut, R. W., and Jones, B., 1997. Controls on aragonite and calcite precipitation in hot spring travertines at Chemurkeu, Lake Bogoria, Kenya. Canadian Journal of Earth Sciences, 34, 801–814. Renaut, R. W., and Jones, B., 2000. Microbial precipitates around continental hot springs and geysers. In Riding, R. E., and Awramik, S. M. (eds.), Microbial Sediments. Berlin: Springer, pp. 187–195. Sheehan, K. B., Patterson, D. J., Dicks, B. L., and Henson, J. M., 2005. Seen and Unseen. Discovering the Microbes of Yellowstone. Guilford, CT: The Globe Pequot Press. Stetter, K. O., 1996. Hyperthermophilic prokaryotes. FEMS Microbiology Reviews, 18, 149–158. Walter, M. R., 1976. Geyserites of Yellowstone National Park: an example of abiogenic stromatolites. In Walter, M. R. (ed.), Stromatolites: Developments in Sedimentology. Amsterdam, Elsevier, pp. 87–112. Waring, G. A., 1965. Thermal springs of the United States and other countries of the world – a summary. United States Geological Survey Professional Paper, 492, 1–383.

Cross-references Extreme Environments Hydrothermal Environments, Fossil Hydrothermal Environments, Terrestrial Microbial Biomineralization Microbial Silicification – Bacteria (or Passive) Sinter

HYDROGEN Tori M. Hoehler NASA-Ames Research Center, Moffett Field, CA, USA

Synonyms Dihydrogen; H2 Definition A diatomic molecule consisting of two atoms of hydrogen or its heavier isotopes. Introduction Hydrogen, the most abundant element in the universe by mass and number, exists on Earth principally in combination with oxygen (in H2O) or carbon (in hydrocarbon and

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organic compounds). The diatomic form, H2 – although present in only trace abundance in the modern oceans and atmosphere – has played a central and evolving role in prebiotic and origin of life chemistry, in linking geochemical and biological cycles, and in shaping the structure, function, and interaction of microbial populations.

Geochemistry of H2 Geosphere H2 is produced by the interaction of water and rocks through a variety of mechanisms (see review in Hoehler, 2005). Among processes that occur in widespread crustal rocks, perhaps the best known and most quantitatively important involves the reduction of water to H2 by the ferrous iron components of rocks, which can be generalized as 2ðFeOÞrock þ H2 O ! ðFe2 O3 Þrock þ H2 ; with (FeO)rock representing the ferrous iron component of igneous rock and (Fe2O3)rock representing the ferric iron component of alteration minerals (e.g., magnetite). The capacity for H2 generation via this mechanism is greatest in minerals with high ferrous iron contents, such as the ultramafic minerals olivine and pyroxene, where the process is called serpentinization. Basalts, with a lower ferrous iron content, have a smaller capacity to produce H2, and granites (where ferrous iron-bearing phases are typically present as minor components) smaller still. The rates of hydrogen generation by water–rock reaction depend significantly on both temperature and the exposure of fresh surface area to water. H2 may also be generated via the radiolytic cleavage of water by a, b, or g radiation produced during decay of isotopes in the host rocks (Spinks and Woods, 1964). Although oxidative species generated simultaneously during the process have potential to recombine with the hydrogen, radiolysis is nonetheless believed responsible for elevated H2 concentrations (up to millimolar levels) in fluid emanations from a number of continental rock units. This mechanism is generally most important for granites, in which the content of the principal radiogenic elements U, Th, and K is generally highest among major crustal rock types. Fracturing of silicate rocks may also serve to liberate H2 that is formed either in solid-state reactions involving trace water impurities in the mineral lattice (Freund et al., 2002) or by direct reduction of water at the fractured surface (Kita et al., 1982). Among these processes, serpentinization offers the greatest potential capacity for H2 generation (per unit volume of rock), by orders of magnitude. However, multiple factors – including mineral composition, fluid flow and chemistry, temperature regime, and mechanical processing – ultimately determine which mode of lithogenic H2 production may prevail in a given environment.

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Atmosphere H2 is a trace component of the modern atmosphere, with an abundance of about 0.5 ppm (Levine, 1985) and a residence time of less than 1 year (Atkinson et al., 1979), but likely higher levels in the early atmosphere would have had important geochemical and biological implications. Atmospheric H2 abundance reflects a balance of sources and sinks. Major source terms (excluding anthropogenic sources) include geological efflux (e.g., volcanic and hydrothermal vent emissions) and biological production. Sinks include oxidative atmospheric reactions, biological consumption, and escape to space. The low abundance of H2 in the modern atmosphere reflects the dominance of oxidative reactions and biological consumption. The early, prebiotic Earth largely lacked such sinks and had a geological source term that was expectedly larger as a result of higher rates of geothermal emanation and a potentially more reducing character in the early mantle and crust (leading to higher H2 concentrations in geothermal emanations). Estimates of atmospheric H2 abundance at this time depend principally on theories regarding the oxidation state of the early mantle and crust, and consideration of the factors constraining H2 escape to space. Even at the low end, these estimates generally predict atmospheric H2 abundance of hundreds to thousands of parts per million, 3–4 orders of magnitude higher than in the present day. H2 itself is considered photochemically inert in the lower atmosphere (Levine, 1985), but it participates in a variety of reactions involving the radical products of photolysis. By serving as a sink for the oxidizing OH radical (Atkinson et al., 1979), and influencing the abundance of the atmospheric reductants H, HCO, and H2CO (Canuto et al., 1983), H2 directly affects the oxidation state of the atmosphere. As such, its relative abundance in the early atmosphere may have significantly affected redox speciation in the volatile nitrogen, carbon, and sulfur systems. In turn, this balance (e.g., the CH4/CO2 ratio, in the carbon system) affects the atmospheric radiation budget (through differing efficiencies of IR absorption), the potential for formation of photochemical hazes (Sagan and Chyba, 1997), and the formation and stability of simple organics in atmospheric chemical processes. Above and beyond its role in buffering the oxidation state of the atmosphere through chemical interactions, escape of hydrogen to space can contribute to the irreversible oxidation of the planet, by carrying electrons out of the system (Walker, 1977). This process occurs at low rates in the modern, oxidized atmosphere, where the low abundance of H2 results in low escape rates. However, H2 escape from an early atmosphere containing orders of magnitude higher H2 concentrations may have contributed substantially to the overall oxidation of the planet. H2 in prebiotic and origin of life chemistry Prebiotic chemistry The emergence of life must depend significantly on development of an inventory of abiotically generated compounds

that can serve as building blocks for biochemistry. Two of the most potentially important endogenous sources for such compounds – synthesis in hydrothermal vent settings and in atmospheric chemical processes – are known or inferred to depend strongly on H2 for the yield and nature of compounds formed. In vent settings, H2 from water–rock reactions is implicated as the reductant in Fischer–Tropsch type synthesis of methane, higher hydrocarbons, and simple organic acids, and is predicted to support the synthesis of amino acids and other prebiotic compounds. Miller–Urey type spark discharge experiments that simulate atmospheric chemical processes have a similar dependence that is demonstrated experimentally. For example, the yield of amino acids from simpler precursors in such experiments depends directly on the abundance of H2, regardless of whether reduced or neutral starting materials (i.e., CH4/NH3 (Miller and Urey, 1959), or CO2/N2 (Schlesinger and Miller, 1983)) are used. Given its significance in prebiotic synthesis and atmospheric redox and photochemistry on the early Earth (see above), H2 likely played an important overall role in establishing the environmental and chemical context from which life emerged.

Origin of life Although the specific chemistry leading to the origin of life is not known, H2 is proposed in some scenarios to have been a key component of the earliest metabolism (Wächtershauser, 1993). The oxidation of H2 serves a potentially important dual purpose with respect to such metabolism, in yielding both reducing power and Hþ. Reducing power is necessary for generating organic biomolecules when the carbon source is CO2. The establishment of Hþ gradients (specifically, across membranes) can be used to store energy for subsequent coupling to biochemistry. Such dual-purpose chemistry underpins the metabolism of a wide range of modern organisms, including those thought to represent Earth’s earliest life. The widespread availability of H2 in geothermal emanations (e.g., Zobell, 1947; Elderfield and Schulz, 1996) and, presumably, in the prebiotic oceans and atmosphere would have made it a ubiquitous and reliable substrate on which to base the first metabolic chemistry. H2 and biology The reversible oxidation of molecular hydrogen, H2 ⇆ Hþ + 2e, is among the most widely utilized reactions in microbial metabolism. Hydrogenases, the enzymes which catalyze this transformation, are present in all three phylogenetic domains of life (Fenchel and Finlay, 1995; Vignais et al., 2001; Schwarz and Friedrich, 2003). They are particularly widely represented in the Bacteria and Archaea, including the most deeply branching and least derived lineages (Vignais et al., 2001; Schwarz and Friedrich, 2003), suggesting that, exclusive of a possible role in origin of life chemistry, H2 metabolism has been an important contributor to biochemistry since the very early stages of evolution.

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Since its emergence in metabolism, H2-based redox chemistry has come to be associated with an extremely broad range of microbial metabolic strategies, including oxygenic and anoxygenic phototrophy, anaerobic chemolitho- and chemoheterotrophy, and aerobic chemolithotrophy (Schwarz and Friedrich, 2003). H2/Hþ can serve as electron donor, acceptor, or both, in these processes. This broad metabolic utility and versatility casts H2 in a highly central role in the ecology of microbial communities, and in the geochemistry they mediate.

Lithotrophic communities Organisms capable of H2 metabolism can potentially be supported directly by geochemical production of H2 (by, e.g., water–rock reaction or radiolysis). As such, they have potential to exist independently of energy input from light or the direct products of photosynthesis, oxygen, and organic matter. Such communities are therefore frequently envisioned as important contributors to Earth’s earliest biosphere, to the rock-hosted subsurface biosphere, and, potentially, to subsurface life on other worlds. H2-based, photosynthesis-independent communities have been documented or inferred for a range of systems and H2-producing mechanisms, including vent fluids sourced in serpentinizing systems, basalt-hosted aquifers, and continental groundwaters bearing a signal of radiolytic H2 production. Heterotrophic communities Systems driven by anaerobic decomposition of organic matter typically host a great diversity of H2-cycling organisms. Under anaerobic conditions, the complete degradation of complex organics cannot be accomplished by a single organism. Instead, the overall process is mediated, in a series of individual steps, by communities of microorganisms having different metabolic capabilities (Schink, 1988). The individual steps are linked through the transfer of electrons from one group of microbes to another, with H2 serving as one of the most commonly employed molecular carriers of electrons. The pool of H2 in such systems is typically characterized by low concentrations (subnanomolar to micromolar) and rapid turnover, with resultant residence times frequently in the range of seconds (Hoehler et al., 2002). Combined with a strong influence on the thermodynamics of reactions that produce or consume it (resulting from its typically high stoichiometric coefficient in such reactions), this effect casts H2 as a key regulator of degradation pathways, community structure, and carbon and energy flow in such environments. Specifically, fluctuations in H2 concentration are documented, in a process known as “interspecies H2 transfer,” to affect the H2-cycling reactions by inhibition or stimulation, alteration of products, or even reversal (see review in Hoehler, 2005). H2 thus occupies a central role in anoxic ecosystems. Anoxia prevailed globally prior to about 2.4 billion years ago and continues to prevail in most organic-bearing

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aqueous environments (including ocean, lake, and wetland sediments and animal digestive tracts). In the past and present, such systems represent a major control on the redox chemistry of the oceans and atmosphere, and the ultimate biological filter on material passing into the rock record.

Phototrophic communities Most of the known photosynthetic microorganisms, representing both anoxygenic and oxygenic photosynthesis, possess a capability to engage in H2 metabolism (Schwarz and Friedrich, 2003; Vignais et al., 1985). Most anoxygenic photosynthesizers, including representatives of the purple sulfur, purple non-sulfur, green sulfur, and green non-sulfur bacteria, can utilize H2 as an electron donor for photosynthetic carbon fixation, and several are also capable of utilizing H2 as an electron donor for the chemical reduction of inorganic oxidants. Some representatives of the cyanobacteria (oxygenic photosynthesizers) are similarly capable when operating in an anoxygenic photosynthetic mode of metabolism. H2 is produced obligately during the conduct of nitrogen fixation and can also be photo-produced by the nitrogenase enzyme complex independently of nitrogen fixation. These capabilities, which occur widely among the cyanobacteria and anoxygenic photosynthesizers, are among the principal mechanisms being explored for biological production of H2 as an alterative energy source. H2 is also generated as a product of fermentation, a capability demonstrated in some cyanobacteria and purple sulfur bacteria. The fermentative mode of H2 production appears to dominate the H2 cycle in a variety of cyanobacterial “mats,” resulting in significant fluxes of H2 out of the system (Hoehler et al., 2001; Hoehler et al., 2002). During the 1–2 billion years of Earth’s history in which cyanobacterial mats dominated biological productivity on Earth, such fluxes may have contributed significantly to atmospheric H2 levels and, by enhanced rates of H2 escape to space, the overall oxidation of the planet. Eukarya Although not as widespread as in the Bacteria and Archaea, H2 metabolism is also found in the Eukarya. In a variety of deep-branching anaerobic Eukaryotes, metabolism is tied to, or dependent on, an H2-based symbiosis with Bacterial or Archaeal partners (see review in Fenchel and Finlay, 1995). The driver of most of these symbioses is an H2generating organelle, the hydrogenosome, which appears in some cases to be derived from a mitochondrion. These hydrogenosomes carry out fermentation, rather than oxidative phosphorylation, of pyruvate – and thereby produce H2. Microbial symbionts (which may include methanogens, sulfate reducers, and (in one case) photosynthetic purple non-sulfur bacteria) increase the energetic yield of this fermentation by consuming the end product H2. The symbiosis significantly enhances the growth of the host, and is, in some cases, so evolved that methanogen symbionts have lost their cell walls, and exhibit synchronization of cell

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division with the host. Among photosynthetic Eukarya, green algae exhibit an active H2 metabolism, and are considered a promising potential source of “biohydrogen” for alternative energy systems.

Summary Hydrogen has played an important role in Earth’s geochemistry and biology since the earliest stages of the planet’s history. H2 is produced by a variety of geochemical processes and is postulated to have been a significant component of the early, prebiotic atmosphere, where it had direct or indirect effects on atmospheric redox chemistry and radiation budget, and contributed to planetary oxidation by escape to space. Prebiotic and origin of life chemistry may have significantly involved H2, and its prominent use among deeply branching organisms in the tree of life reflects, at very least, a key role in metabolism since the biosphere’s early stages. In the modern biosphere, H2 biochemistry is associated with a broad variety of metabolic strategies, in all three domains of life. Although it now represents a trace component of the oceans and atmosphere, the significance of H2 for planetary chemistry persists through its central role in the microbial world. Bibliography Atkinson, R., Darnall, K. R., Lloyd, A. C., Winer, A. M., and Pitts, J. N. Jr., 1979. Kinetics and mechanisms of the reaction of the hydroxyl radical with organic compounds in the gas phase. Advances in Photochemistry, 11, 375–488. Canuto, V. M., Levine, J. S., Augustsson, T. R., Imhoff, C. L., and Giampapa, M. S., 1983. The young sun and the atmosphere and photochemistry of the early Earth. Nature, 305, 281–286. Elderfield, H., and Schulz, A., 1996. Mid-ocean ridge hydrothermal fluxes and the chemical composition of the ocean. Annual Review of Earth Planetary Science, 24, 191–224. Fenchel, T., and Finlay, B. J., 1995. Ecology and Evolution in Anoxic Worlds. Oxford: Oxford University Press. Freund, F., Dickinson, J. T., and Cash, M., 2002. Hydrogen in rocks: an energy source for deep microbial communities. Astrobiology, 2, 83–92. Hoehler, T. M., 2005. Biogeochemistry of dihydrogen. In Sigel, A., Sigel, H., and Sigel, R. K. O. (eds.), Biogeochemical Cycles of Elements. New York: Marcel Dekker, Vol. 43, pp. 9–48. Hoehler, T. M., Bebout, B. M., and DesMarais, D. J., 2001. The role of microbial mats in the production of reduced gases on the early Earth. Nature, 412, 324–327. Hoehler, T. M., Albert, D. B., Alperin, M. J., Bebout, B. M., Martens, C. S., and DesMarais, D. J., 2002. Comparative ecology of H2 cycling in sedimentary and phototrophic ecosystems. Antonie Van Leeuwenhoek, 81, 575–585. Kita, I., Masuo, S., and Wakita, A., 1982. H2 generation by reaction between H2O and crushed rock: an experimental study of H2 degassing from the active fault zone. Journal of Geophysical Research, 87, 10789–10795. Levine, J. S., 1985. The photochemistry of the early atmosphere. In Levine, J. S. (ed.), The Photochemistry of Atmospheres. San Diego, CA: Academic Press, p. 518. Miller, S. L., and Urey, H., 1959. Organic compound synthesis on the primitive Earth. Science, 130, 245–251. Sagan, C., and Chyba, C., 1997. The early faint sun paradox: organic shielding of ultraviolet-labile greenhouse gases. Science, 276, 1217–1221.

Schink, B., 1988. Principles and limits of anaerobic degradation. In Zehnder, A. J. B. (ed.), Biology of Anaerobic Microorganisms. New York: Wiley-Interscience, pp. 771–846. Schlesinger, G., and Miller, S. L., 1983. Prebiotic synthesis in atmospheres containing CH4, CO, and CO2: II. Hydrogen cyanide, formaldehyde, and ammonia. Journal of Molecular Evolution, 19, 383–390. Schwarz, E., and Friedrich, B., 2003. The H2-metabolizing prokaryotes. In Dworkin, M., et al. (eds.), The Prokaryotes: An Evolving Electronic Resource for the Microbiological Community. New York: Springer. Spinks, J. W. T., and Woods, R. J., 1964. An Introduction to Radiation Chemistry. New York: Wiley. Vignais, P., Colbeau, A., Willison, J. C., and Jouanneau, Y., 1985. Hydrogenase, nitrogenase, and hydrogen metabolism in the photosynthetic bacteria. In Rose, A. H., and Tempest, D. W. (eds.), Advances in Microbial Physiology. New York: Academic Press, Vol. 26, pp. 155–234. Vignais, P. M., Billoud, B., and Meyer, J., 2001. Classification and phylogeny of hydrogenases. FEMS Microbiology Reviews, 25, 455–501. Wächtershauser, G., 1993. The cradle chemistry of life: on the origin of natural products in a pyrite-pulled chemo-autotrophic origin of life. Pure and Applied Chemistry, 65(6), 1343. Walker, J. C. G., 1977. Evolution of the Atmosphere. New York: Macmillan. Zobell, C. E., 1947. Microbial transformation of molecular hydrogen in marine sediments, with particular reference to petroleum. Bulletin of the American Association of Petroleum Geochemists, 31(10), 1709–1751.

Cross-references Acetogens Anaerobic Oxidation of Methane with Sulfate Astrobiology Chemolithotrophy Early Earth Fermentation Hydrothermal Environments, Terrestrial Microbial Communities, Structure, and Function Microbial Degradation Origin of Life Photosynthesis Sulfate-Reducing Bacteria Symbiosis Terrestrial Deep Biosphere

HYDROTHERMAL ENVIRONMENTS, FOSSIL Joachim Reitner University of Göttingen, Göttingen, Germany

Synonyms Black and White Smoker communities; Volcanogenic massive sulfide deposits – VMS Definition “Black smokers” represent extreme hydrothermal activities that are commonly located in deep sea environments

HYDROTHERMAL ENVIRONMENTS, FOSSIL

related with mid-ocean ridges and/or oceanic crust fault systems. Hydrothermal vent systems result from hydrothermal circulation of fluids controlling the transfer of energy and various matter from the interior to the surface of the oceanic crust, respectively the sediment surface. This hydrothermal circulation is a fundamental geological process and has been permanently influencing the chemistry of the oceans, and in consequence, also the composition of the entire earth crust. Seafloor hydrothermal activity has a major impact on the chemistry and life processes of the oceans (Parson et al., 1995; Humphries et al., 1995; van Dover, 2000). This entry focuses on volcanogenic massive sulfide deposits (VMS) and not on sedimentary exhalative deposits (SEDEX) – however, the boundary between both environments is not very strict. Massive sulfide deposits, hydrothermally altered rocks, and special communities of organisms are an important record of this process. Ancient seafloor hydrothermal activity is recorded in massive sulfides from the 3.5 GY-old Pilbara region of Western Australia (Barley, 1992). The new findings of hydrothermal vents with microbialites from 1.43 GY-old Precambrian rocks in northern China (Li and Kusky, 2007) are also of great interest. Within the Phanerozoic, roughly 20 sites are known with fossil hydrothermal vent communities (Little et al., 1998).

Geobiological notes Hydrothermal vents have become one of the most fascinating fields of research in marine sciences. The recognition of complex vent communities consisting of organisms dependent on chemolithotrophy and extreme conditions of very high temperature, pressure, and chemistry at the ocean ridges was a great discovery (Humphris et al., 1995). There are many reasons to believe that life originated at hydrothermal vent environments in the late Hadean. However, only few publications are dealing with the fossil record of hydrothermal environments connected with preserved organism communities. There are about 20 known Phanerozoic VMS sites with preserved hydrothermal vent communities, ranging from the Silurian to the Eocene. Most of these sites are found in the Ural Mountains and on Cyprus, some of them in Ireland, California, Philippines, and New Caledonia. All occurrences are linked to remains of oceanic crust (ophiolites). The investigation of vent systems is still a challenge, because the sites are normally small and most of them were destroyed during subduction of oceanic crust. The vent communities contain assemblages of inarticulate and rhynchonellid brachiopods, gastropods, bivalves, and also monoplacophorans (Little et al., 1998; Kiel and Little, 2006). Remains of worm tubes are very characteristic and may be related to alvinellid polychaetes and pogonophorid-vestimentiferan chemotrophic worms. Most of the fossil taxa of the vent sites are endemic. However, the diagenetic loss of the original shell material makes it very difficult to classify the worm tubes in a modern

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taxonomic scheme. Surprisingly, no arthropods were found at the fossil sites, whereas arthropods are very common in modern vent environments. There is an ongoing discussion on the origin of the vent fauna. New groups of organisms may have immigrated from the ambient environment and adapted to the extreme conditions of the vent sites and vice versa, some of them have left this environment. The origin of the vent faunas is thoroughly discussed in Kiel and Little (2006) and Little and Vrijenhoek (2003). Kiel and Little (2006) suggested that the mollusk seep faunas (hydrothermal vents and cold seeps) have a significantly longer evolutionary history than normal marine mollusks and show a tight taxonomic relationship with deep-sea mollusk faunas. The oldest known Phanerozoic hydrothermal vent is of Silurian age (Little et al., 1998). The discovery of the Mesoproterozoic (1.43 GY) hydrothermal vents in northern China was of great importance due to very wellpreserved filamentous microbial communities forming massive sulfidic microbialites (Li and Kusky, 2007).

Conclusion Fossil hydrothermal vent systems are known since the early Archaean and represent important geobiological environments. These environments have a significant key role in biological evolutionary processes. However, only few (ca. 20) sites are known worldwide in the Phanerozoic with preserved vents faunas. The oldest known fossil hydrothermal vent system with a metazoan community is known from the Silurian of the Ural Mountains (Little et al., 1998). Many of the vent organisms probably have a significantly longer evolutionary history than normal marine taxa (Kiel and Little, 2006). Bibliography Barley, M. E., 1992. A review of Archaean volcanic-hosted massive sulphide and sulphate mineralisation in Western Australia. Economic Geology and the Bulletin of the Society of Economic Geologists, 87, 855–872. Humphris, S. E., Zierenberg, R. A., Mullineaux, L. S., and Thomson, R. E., 1995. Seafloor hydrothermal systems. Geophysical Monograph (AGU), 91, 1–466. Kiel, S., and Little, C. T. S., 2006. Cold seep mollusks are older than the general marine mollusk fauna. Science, 313, 1429–1431. Li, J., and Kusky, T. M., 2007. World’s largest known Precambrian fossil black smoker chimneys and associated microbial vent communities, North China: implications for early life. Gondwana Research, 12, 84–100. Little, C. T. S., and Vrijenhoek, R. C., 2003. Are hydrothermal vent animals living fossils? Trends in Ecology and Evolution, 18, 582–588. Little, C. T. S., Herrington, R. J., Maslennikov, V. V., and Zaykov, V. V., 1998. The fossil record of hydrothermal vent communities. In Mills, R. and Harrison, K. (eds.), Modern Ocean Floor Processes and the Geological Record. Geological Society London Special Publications, 148, pp. 259–270. Parson, L. M., Walker, C. L., and Dixon, D. R., 1995. Hydrothermal vents and processes. Geological Society London Special Publication, 87, 1–411. Van Dover, C. L., 2000. The Ecology of Deep-Sea Hydrothermal Vents. Princeton, NJ: Princeton University Press, 424 p.

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Cross-references Cold Seeps Deep Biosphere of the Ocean Deep Sea Extreme Environments Hydrothermal Environments, Marine Iron Sulfide Formation Origin of Life

HYDROTHERMAL ENVIRONMENTS, MARINE Gilberto E. Flores, Anna-Louise Reysenbach Portland State University, Portland, OR, USA

Definition Marine hydrothermal environments are one of the most extreme environments on Earth, yet they support highly productive biological communities over a wide range of physical and chemical environments. The following review is a general discussion of the types of microorganisms found in these different environments preceded by brief descriptions on hydrothermal fluid generation, mineral deposit formation, and microbial metabolism. Introduction The discovery of deep-sea hydrothermal vents and the lush biological communities associated with them in the late 1970s ushered in a new era for marine biologists. Prior to this discovery, life in the deep-sea was thought to be dependent on the settling of detrital material from the productive, overlying surface waters. While this is true for much of the ocean basins, the level of production observed at these newly discovered vent sites was too great to be supported by these mechanisms alone. It soon became evident that life in these environments was supported by chemosynthetic primary production in which microorganisms harnessed the abundant geochemical energy available in the hydrothermal fluids. This challenged one of the basic ecological premises that all ecosystems on Earth were dependent on light energy and driven by photosynthetic primary production. Since those initial revelations, research efforts have focused on assessing the microbial diversity and attempting to understand the intimate associations between microbial productivity, geology, and geochemistry in marine hydrothermal environments. This article is a general discussion of the trends that are beginning to emerge from these efforts. Included in the discussion are overviews of the geologic setting of marine hydrothermal systems, the generation of hydrothermal fluids from different geologic settings, and the types of structures formed. For additional in-depth reviews, the Geophysical Monographs published by the American Geophysical Union (AGU) are valuable resources (Humphris et al.,

1995; German et al., 2004; Wilcock et al., 2004; Christie et al., 2006).

Geological setting Most seafloor hydrothermal vent systems are associated with extensional tectonic activity and heated by magmatic heat as it is convected into the crust (Seyfried and Mottl, 1995). The most well-studied areas of hydrothermal circulation are along divergent plate boundaries where basaltic seafloor is made and mid-ocean ridges (MOR) arise (Figure 1). Hydrothermal circulation along MORs results from the active heating of seawater that percolates through newly formed basaltic crust. Hydrothermal venting is also commonly found along convergent plate margins where an oceanic plate is subducted beneath a continental plate forming island arcs and back-arc basins (Christie and Fisher, 2006). Vents along back-arc basins are actively heated in the same way as those along MOR, although the fluids tend to be more heterogeneous due to the variability in magma composition and additional inputs from the subducting plate (GAOM et al., 2006). A third location where seafloor hydrothermal systems can be found is at intraplate volcanic hot spots. Volcanic hot spot systems are not directly associated with tectonic plate margins, but are actively heated as plumes of molten magma push up through the mantle and crust (Seyfried and Mottl, 1995). The recently discovered Lost City vent field more than 15 km off axis of the Mid-Atlantic Ridge (MAR) represents a new type of seafloor hydrothermal system as it appears to be at least partially heated by exothermic reactions between mantle derived peridotite and seawater (Kelley et al., 2001). These reactions have a dramatic impact on the composition of the hydrothermal fluids, which in turn affects the microbial community as discussed in the following sections. Although Lost City represents the first vent type of its kind discovered, similar systems are believed to exist along margins of slow- and ultraslow-spreading ridges where uplifting of ultramafic massifs are common. Hydrothermal fluids Seawater heated in contact with subsurface rocks dramatically alters the chemical composition of the end member fluids. The degree of alteration is influenced by several factors including the initial composition of the seawater, the type and structure of the host rock, the presence of sediment overlaying the host rock, and the type, depth, and size of the heat source (reviewed in Tivey, 2007). Understanding these interactions and how they affect the composition of hydrothermal fluids is important as the fluids set the physiological parameters and provide the metabolic menu for colonizing microbes. For comparison, representative compositions of hydrothermal fluids from different settings are presented in Table 1.

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Hydrothermal Environments, Marine, Figure 1 Global distribution of known (red) hydrothermal venting along mid-ocean ridges (MOR), back-arc basins, rifted arcs, and at submerged island-arc volcanoes. Yellow circles indicate areas where mid-water column chemical anomalies have been detected suggesting hydrothermal activity. EPR East Pacific Rise; TAG Trans Atlantic Geotraverse; MEF Main Endeavor Field; GR-14 Sea Cliff hydrothermal field on the northern Gorda Ridge. Figure 1 from Tivey (2007).

Basalt hosted hydrothermal fluids Water–rock reactions begin as cold seawater infiltrates shallow basalt in the recharge zones (Figure 2). Initial low temperature reactions (60 C) result in the removal of alkali metals from seawater and the oxidation of basaltic minerals (i.e., olivine and plagioclase), with the concomitant leaching of silica (Si), sulfur (S), and sometimes magnesium (Mg) from the minerals (reviewed in Alt, 1995). Magnesium is eventually completely removed from the fluids as Mg-rich clays precipitate at temperatures above 150 C. Anhydrite (CaSO4) precipitation and seawater sulfate (SO42) reduction also occur in the recharge zone at temperatures above 150 C, resulting in the complete removal of SO42 from the fluids. As the fluid continues through the crust, reactions with ironbearing minerals such as olivine and pyroxene result in highly reducing fluids with elevated concentrations of hydrogen gas (H2) (Tivey, 2007). Upon reaching the reaction zone where fluids are heated upward of 400 C, the fluid obtains its chemical signature by leaching S and metals (copper-Cu, iron-Fe, manganese-Mn, zinc-Zn) from the surrounding rocks (Alt, 1995). Phase separation

may also occur in the reaction zone if temperature and pressure are greater than 407 C and 298 bars, respectively (Von Damm, 1995). Phase separation can have a significant impact on the composition of end member fluids as volatiles such as hydrogen-sulfide (H2S) tend to partition in the vapor phase while metals such as Fe become enriched in the liquid (brine) phase. Gases such as helium (He), methane (CH4), and carbon dioxide (CO2) may also be added to the hydrothermal fluids as volatiles from the underlying magma (Alt, 1995). In total, these reactions result in end member fluids that are acidic (pH 2.8–4.5), highly reduced, rich in H2S and metals, and heated up to 400 C. The fluid is also extremely buoyant relative to seawater and is forced back to the seafloor through the discharge zone and may emerge as diffuse or focused flow depending on the degree of subsurface mixing during the ascent (Figure 2). Other factors, such as differences in host rock and sediment cover, can contribute to the chemical composition of end member fluids. The impact that different host rocks have on fluid chemistry is best illustrated by the ultramafic hosted Rainbow vent field along the MAR

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Hydrothermal Environments, Marine, Table 1 Representative chemical compositions of hydrothermal fluids from different geologic settings Characteristic

Deep seawater

Mid-ocean ridge

Back-arc basin

Sediment hosted

Rainbow vent field

Lost City

T ( C) pH (25 C) SO4, mmol/kg Mg, mmol/kg Cl, mmol/kg Na, mmol/kg Ca, mmol/kg K, mmol/kg H2S, mmol/kg H2, mmol/kg CO2, mmol/kg CH4, mmol/kg NH3, mmol/kg Fe, mmol/kg Mn, mmol/kg Cu, mmol/kg Zn, mmol/kg Pb, mmol/kg

2 8 28 53 545 464 10.2 10.1 – – 2.36 – – – – – – –

405 2.8–4.5 0 0 30.5–1,245 10.6–983 4.02–109 1.17–58.7 0–19.5 0.0005–38 3.56–39.9 0.007–2.58 60% up to 3000 rel.1/cm when using a 532 nm laser. Using the same grating with a 785 nm laser will cause the efficiency to drop to about 30% at 3000 rel.1/cm. The correct grating blazed at 750 nm, however, shows an efficiency >60% up to 3000 rel.1/cm when used with the 785 nm laser. Efficiency of the spectrometer: There are a variety of spectrometers available on the market. While mirror-based spectrometers can generally be used over a wide spectral range, lens-based spectrometers often show a higher throughput in the spectral range they are designed for. Some lens-based spectrometers show a transmission at 532 nm of >60%. Spectrometers using a combination of lenses and prisms or mirrors generaly have a significantly lower throughput. Commercially available mirror-based

Czerny-Turner spectrometers only have a transmission of about 30% at this wavelength. Efficiency of the detector (CCD): Back-illuminated CCD cameras show a quantum efficiency (QE) of >90% over the entire spectral range of interest for 532 nm excitation. Deep depletion back-illuminated CCD cameras, on the other hand, show a QE of >90% for 785 nm excitation. As a comparison, front-illuminated CCD cameras do not exceed 55% quantum efficiency at any wavelength. Additionally, the dark current of the cameras needs to be minimized, which is achieved through efficient Peltier cooling. The readout noise is another limiting factor for small signals. As the analog to digital (A/D) converter of all cameras will add at least 5–10 electrons read-out noise to the signal, any signal below approximately 5 electrons will be lost in the noise. Additionally, the faster the A/D converter is operating, the higher the read-out noise will be. Electron-multiplying CCD (EM-CCD) cameras can be used to overcome this problem. With these cameras, the signal is amplified before the A/D conversion, allowing the detection of even single photons and reducing the necessary integration time down to milliseconds. Principle of operation Confocal Raman microscopes generally provide a variety of modes of operations. The most common are listed below: Collection of Raman spectra at selected sample areas (single spectrum): Single Raman spectra can be collected at user-selectable sample areas with integration times ranging from ms to hours. The position of the collected spectrum can normally be fully controlled in 3D. A stable and precise positioning system must be included in the instrument to ensure that the point of interest will remain fixed under the excitation focus. This is very important when spectra with longer integration times for the best quality and signal to noise ratio are to be obtained from extremely small sample volumes. For example, using an oil immersion objective (NA 1.4) with a 532 nm laser and the proper pinhole size allows the sample volume to be as small as 230  230  550 nm. Collection of time series of Raman spectra at selected sample areas (time spectrum): With this mode, time series of Raman spectra can be obtained to analyze dynamic sample properties. Thousands of spectra can be obtained over time and analyzed with integration times ranging from ms to tens of seconds. Raman spectral imaging: In the Raman spectral imaging mode, the sample is moved in X and Y and a full Raman spectrum is obtained at every pixel measured. From these data sets images of, e.g., the integrated intensity of various bands can be generated. This is illustrated in the following example. A drill core section from the Äspö Hard Rock Laboratory, Sweden, was studied using the large area scan mode CRM, aiming to characterize secondary fracture fillings in

RAMAN MICROSCOPY (CONFOCAL)

a 1.8–1.4 billion years old diorite. The rock sample was obtained from 450 m below the surface (kindly provided by SKB, Swedish nuclear fuel and waste management). This sample was examined with a 532 nm laser and a 50 air objective (NA 0.55). The scan range was 8000 μm in X and 2000 μm in Y with 800  200 points resolution. Each spectrum was integrated for 36 ms. Figure 6 shows the characteristic spectra of calcite, fluorite and quartz found in the sample. Integrating over, for example, the area marked in green in Figure 6d results in a single value for each of the 160,000 spectra. These spectra can be displayed as a color-coded image as shown in Figure 6a for quartz. Here, brighter values indicate a higher integrated intensity of the quartz peak and the distribution of quartz can thus be obtained from this image.

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Other spectra of the same scan show the characteristic features of calcite or fluorite (Figure 6d). Using these, the calcite (Figure 6b) and fluorite (Figure 6c) images can be generated by using additional integral filters for the marked regions. Other features of the spectra such as the width of peaks or their position can also be evaluated by applying the corresponding filters. Further evaluation of the data allows the averaging of similar spectra (for which a cluster analysis is often used) and the subtraction of, for example, pure spectra from mixed spectra to extract the spectra of the various components (spectral de-mixing). These spectra can then be used with the basis analysis, where each of the spectra recorded are fitted with a linear combination of the basis spectra. The result of such an analysis is one image for each basis

Raman Microscopy (Confocal), Figure 6 Spectra (d) and spectral images of quartz (a), calcite (b) and fluorite (c) recorded from the diorite (a) and the fracture fillings (b and c). The data were recorded with a WITec alpha500R confocal Raman microscope.

Raman Microscopy (Confocal), Figure 7 DE-mixed and averaged spectra of the various components in the diorite and the adjacent fracture minerals with the combined image on the left.

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Raman Microscopy (Confocal), Figure 8 Variations in the quartz phases due to changes in the crystallographic orientation, the crystallinity, and the minerals.

spectra which can then be combined to generate a false color image showing the distribution of all components (Figure 7). The color-coded Raman image and corresponding spectra in Figure 7 allow for the general assignment of mineral phases and their gross distribution over the scanned area. In addition to the mineralogical context information, organic components were identified, spectrally characterized and located, trapped between two generations of fracture fillings (the hydrothermal fluorite and low temperature calcite, Tullborg et al., 2008; Wallin and Petermann, 1999). This information allowed to infer at which point in time a “deep biosphere” (see entry “Terrestrial Deep Biosphere”) was active within these rocks. Cluster analysis of the data set revealed discrete areas of variation in the mineral phases (Figure 8). This is exemplified by the quartz phase where four distinct regions were identified based on variations in relative peak intensity. Plotting regions of equal intensities of the quartz line at ca. 200 cm1 shows discrete regions in the sample corresponding to each of the identified spectra. Quartz was selected as an example to highlight the feasibility of color-coded Raman imaging to locate changes of different mineral phases. These changes can likely be attributed to different levels of crystallinity and crystal orientation. It is noteworthy that the discrete phase colored green only occurs at the interface with the plagioclase minerals, which can be attributed to a secondary phase due to the alteration and phase changes of the plagioclases. Since SiO2-phases play an important role in biomineraliation, this example highlights the potential benefits confocal Raman imaging may provide in understanding the processes and dynamics involved in these processes.

Summary CRM is a nondestructive analytical technique that merges Raman spectroscopy and confocal microscopy for the visualization of molecular information over a defined sample area. The technique makes use of the Raman effect (Raman, 1928), i.e., the energy shift between exciting and scattered photons which is caused by the excitation (or annihilation) of a molecular vibration. This energy shift is characteristic and therefore a fingerprint for the type and coordination of the molecules involved. By means of CRM, the spatial distribution and association of components in the sample, including organics as well as minerals, can be evaluated from large scale scans in the centimeter range to the finest detail with submicron resolution. This way, CRM may contribute significantly to the understanding of a sample’s chemical composition, and complexity, in geological and geobiological studies. Bibliography Fries, M., and Steele, A., 2010. Raman spectroscopy and confocal Raman imaging in mineralogy and petrography. In Dieing, T., Hollricher, O., and Toporski, J. (eds.), Confocal Raman Microscopy, Heidelberg: Springer. Hild, S., Marti, O., and Ziegler, A., 2008. Spatial distribution of calcite and amorphous calcium carbonate in the cuticle of the terrestrial crustaceans Porcellio scaber and Armadillidium vulgare. Journal of Structural Biology, 142, 100–108. Ibach, H., and Lüth, H., 2003. Solid State Physics. An Introduction to Principles of Materials Science, Berlin: Springer. Raman, C., 1928. A new radiation. Indian Journal of Physics, 2, 387. Raman, C., and Krishnan, K., 1928. A new type of secondary radiation. Nature, 121, 501. Smekal, A. G., 1923. Zur Quantentheorie der Dispersion, Naturwissenschaften, 11, 873–875.

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Tullborg, E.-L., Drake, H., Sandström, B., 2008. Palaeohydrogeology: A method based on fracture mineral studies. Applied Geochemistry, 23, 1881–1897. Wallin, B., and Peterman, Z., 1999. Calcite fracture fillings as indicators of palaeohydrology at Laxemar at the Äspö Hard Rock Laboratory, southern Sweden. Applied Chemistry, 14, 953–962. Wilson, T., 1990. Confocal Microscopy, London: Academic.

Cross-references Biosignatures in Rocks Terrestrial Deep Biosphere

REDUCTION SPHEROIDS Beda A. Hofmann Natural History Museum Bern, Bern, Switzerland

Synonyms Bleaching haloes; Fish eyes; Radioactive concretions; Radioactive nodules; Reduction mottling; Reduction spots; Reduktionshöfe Definition Reduction spheroids are small-scale (diameter: few millimeters to maximum about 20 cm) spheroidal bleached features in red-bed sediments characterized by the absence of hematite and often containing a mineralized core (center, Figure 1). Reduction spheroids result from local chemical reduction of ferric iron and other elements. Host rocks are marine and continental red beds and, more rarely, altered, hematite-stained crystalline rocks. Reduction spheroids have been described from many occurrences in red beds of widely variable age worldwide: Proterozoic of Canada (Tanton, 1948); Cambro-Silurian marine red beds of the NE USA (Beutner and Charles, 1985); Devonian continental red beds of Scotland (Parnell, 1985); Carboniferous continental red beds of New Brunswick and Nova Scotia, Canada (Dyck and McCorkell, 1983); Permian continental red beds of Germany (Eichhoff and Reineck, 1952; Mempel, 1960; Schreiter, 1930), Switzerland (Hofmann, 1990, 1991), Southwest England (Harrison, 1970; Perutz, 1940), and Cretaceous radiolarian cherts of Oman (Hofmann, 1991). Reduction spheroids characteristically consist of a spheroidal volume of rock from which hematite pigment has been dissolved. Central dark cores often display pronounced concentric zonation. The cores are strongly enriched in numerous redox-sensitive elements (dominant V, but also U, Se, As, Ni, Co, Cu, Ag, Au, PGE, and others). The dominant mineral in reduction spheroid cores is the vanadium mica roscoelite KV2AlSi3O10(OH)2. Uraninite is another very common mineral, leading to an often increased radioactivity of reduction spheroids (Figure 2). Arsenides of Ni, Co, and Cu are also quite common. Radiometric dating of roscoelite, uraninite, and

Reduction Spheroids, Figure 1 Reduction spheroids in Triassic red bed (marly dolomite), Chinle Formation, Paradox Valley, Colorado, USA. These reduction spheroids are strongly enriched in V, Cr, and Au. Diameter of larger reduction spheroid is 8 cm.

Pb-depleted haloes indicates late diagenetic ages (Hofmann, 1990; Hofmann and Frei, 1996). Reduction spheroids often occur in large numbers in red beds and can constitute up to several vol% of the rock. Based on textural evidence, such as fracture-bound occurrences and mass-balance calculations (Hofmann, 1991), reduction spheroids cannot be due to detrital organic-rich particles. Instead, the formation must be due to locally catalyzed redox reactions between a mobile reductant and oxidants. The presence of isotopically light sulfides in some cores (Hofmann, 1991), indicating diagenetic sulfate reduction, as well as the need for a local catalyst, strongly favors a microbiological origin of reduction spheroids. Reductants may be mobile hydrocarbons or molecular hydrogen resulting from the radiolysis of pore water (Hofmann, 1992). The formation of reduction spots is most likely due to chemolithotrophic communities. Kerogen-like organic matter is occasionally present at some localities (Curiale et al., 1983; Parnell, 1985) but is lacking at many other occurrences (Hofmann, 1993).

Conclusion Reduction spheroids are very common in red-bed sediments, and there are strong indications for an origin due to redox reactions between a mobile reductant and oxidized elements locally catalyzed by microbial activity. Reduction spheroids are thus easily visible features resulting from redox processes catalyzed by a deep biosphere of sediments, and represent a type of biosignature that may also be present in oxidized rocks on Mars. Reduction spheroids contain small-scale

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REEFS

Reduction Spheroids, Figure 2 a-Autoradiographs of uranium-enriched reduction spheroid cores, Weiherfeld well, Permian of northern Switzerland. The accumulation of uranium reflects the heterogeneous distribution of reduction activity, consistent with microbial activity. Diameter of radioactive uranium-rich zone is (a) 5 mm and (b) 10 mm.

element enrichments that are in many ways analogous in element inventory, geochemistry, and mineralogy to ore deposits such as unconformity-related and roll-type uranium ores.

Bibliography Beutner, E. C., and Charles, E. G., 1985. Large volume loss during cleavage formation, Hamburg sequence, Pennsylvania. Geology, 13, 803–805. Curiale, J. A., Bloch, S., Rafalska-Bloch, J., and Harrison, W. E., 1983. Petroleum-related origin for uraniferous organic-rich nodules of southwestern Oklahoma. AAPG Bulletin, 67, 588–608. Dyck, W., and McCorkell, R. H., 1983. A study of uranium-rich reduction spheroids in sandstones from Pugwash Harbour, Nova Scotia. Canadian Journal of Earth Sciences, 20, 1738–1746. Eichhoff, H. J., and Reineck, H. E., 1952. Uran-Vanadinkerne mit Verfärbungshöfen in Gesteinen. Neues Jahrbuch für Mineralogie, Monatshefte, 11/12, 294–314. Harrison, R. K., 1970. Hydrocarbon-bearing nodules from Heysham, Lancashire. Geological Journal, 7, 101–110. Hofmann, B. A., 1990. Reduction spheroids from northern Switzerland: mineralogy, geochemistry and genetic models. Chemical Geology, 81, 55–81. Hofmann, B. A., 1991. Mineralogy and geochemistry of reduction spheroids in red beds. Mineralogy and Petrology, 44, 107–124. Hofmann, B. A., 1992. Isolated reduction phenomena in red beds: a result of porewater radiolysis? In Maest A. S. (ed.), 7th International Symposium on Water-Rock interaction, Balkema, Park City, Utah, USA, pp. 503–506. Hofmann, B. A., 1993. Organic matter associated with mineralized reduction spots in red beds. In Parnell, J., Kucha, H., and Landais, P. (eds.), Bitumens in Ore Deposits. Berlin: Springer, pp. 362–378. Hofmann, B. A., and Frei, R., 1996. Age constraints of reduction spot formation from Permian red bed sediments, northern Switzerland, inferred from U-Th-Pb systematics. Schweizerische Mineralogische und Petrographische Mitteilungen, 76, 235–244. Mempel, G., 1960. Neue Funde von Uran-Vanadiumkernen mit Entfärbungshöfen. Geologische Rundschau, 49, 263–276.

Parnell, J., 1985. Uranium/rare earth-enriched hydrocarbons in Devonian sandstones, northern Scotland. Neues Jahrbuch für Mineralogie—Monatshefte, H3, 132–144. Perutz, M., 1940. Radioactive nodules from Devonshire, England. Mineralogische und Petrographische Mitteilungen, 51, 141–161. Schreiter, R., 1930. Vanadinhaltige Kerne, Bleichungsringe und Bleichungszonen in den Schieferletten des Rotliegenden von Sachsen. Zeitschrift der Deutschen Geologischen Gesellschaft, 82, 41–47. Tanton, T. L., 1948. Radioactive nodules in sediments of the Sibley Series, Nipigon, Ontario. Transactions of the Royal Society of Canada, 17(Ser. III, Sec. 4), 69–75.

Cross-references Biosignatures in Rocks Chemolithotrophy Deep Biosphere of Sediments Hydrogen Ores, Microbial Precipitation and Oxidation Radioactivity (Natural)

REEFS Reefs are laterally confined submarine carbonate structures developed by the growth or activity of sessile benthic aquatic organisms, such as corals, bryozoans, sponges, algae, and microbes (Flügel and Kiessling, 2002). See entry “Carbonate Environments” for further reading and classification.

Bibliography Flügel, E., and Kiessling, W., 2002. A new look at ancient reefs. In Kiessling, W., Flügel, E., and Golonka, J. (eds.), Phanerozoic Reef Patterns. Tulsa, OK: SEPM Special Publication Series No. 72, pp. 3–20.

RNA-WORLD

REMINERALIZATION (OF ORGANIC MATTER) Degradation of organic matter that results in net conversion of organic carbon to inorganic (oxidized) carbon species such as CO2 or HCO3-. See entries “Carbon (Organic, Cycling)” and “Carbon (Organic, Degradation)” for further reading.

RHODOPHYTA The Rhodophyta, also referred to as “red algae,” are a distinct eukaryotic lineage of algae with simple plastids (two membranes, derived from primary endosymbiosis). Rhodophytes have a long geological record since the Mesoproterozoic (1.2 Ga). Today, they encompass about 670 largely marine genera with up to 4,500 species predominating the coastal and continental shelf areas of tropical, temperate, and cold-water regions. Red algae are significant as primary producers and providers of structural habitat for other marine organisms. Calcifying (coralline) red algae play an important role in the primary establishment and maintenance of limestone reefs. For further details, see entry “Algae (Eukaryotic).”

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Cross-references Algae (Eukaryotic) Carbonate Environments Symbiosis

RNA-WORLD The term “RNA-World,” first suggested by Walter Gilbert in 1986, describes a popular concept on a hypothetical early stage in the origin of life on Earth. Based on discoveries of enzymatic activities in microbial ribonucleic acids (RNA), a system was suggested where primordial RNA molecules accounted for both, the storage of genetic information and catalytic functions necessary to assemble themselves in a primitive self-replicating system. The RNA molecules evolved in self-replicating patterns, using recombination and mutation to explore new functions and to adapt to new niches (Gilbert, 1986). Such a system would not have required the presence of DNA and/or protein enzymes for biochemical reactions. For further reading, please refer to “Origin of Life.”

Bibliography Gilbert, W., 1986. The RNA world. Nature, 319, 618.

S

SALINE LAKES Carol D. Litchfield George Mason University, Manassas, VA, USA

Synonyms Salt Lakes Definition Salt: chloride, sulfate, phosphate, carbonate, bicarbonate salts of primarily sodium, magnesium, calcium, and potassium. Lake: an enclosed body with or without water entering it from a stream, rain, or snow melt. Ephemeral lake: a lake formed only following rain or snow melt, which dries up during the rest of the year. Thalassohaline: salts in a lake derived from the evaporation of seawater with the dominant ions of sodium and chloride as found in the ocean. Athalassolhaline: salts in a lake derived from rocks and geological weathering and dominated by magnesium, calcium, and sulfate. Alkaliphilic lake: also called soda lake. A lake in which the dominant ions are sodium and bicarbonate and carbonate. This causes the lake to have an alkaline pH. General description of saline lakes Saline lakes are a natural worldwide phenomenon. They can be found on all continents and in most countries, and they are frequently recognizable by their name – for example, Salt Lick, Salt Lake, Salt Pond, and Salt Marsh. They range in salinity from just above the salt content of sea water (>3.2% salts) to hypersaline lakes (>20% and up to saturation with respect to salts). Examples of saline lakes include, at the lower end, salt lakes in Antarctica, the Caspian Sea, which borders five countries and has

a salinity slightly less than that of seawater, and the Salton Sea in California. At the upper end in salt concentrations are the Dead Sea bordering Israel and Jordan, the soda lakes of North Africa, and Quinghai Lake in China. A general characteristic of saline lakes is that they are terminal lakes, which means that although rivers may flow into them, they have no significant outflow. Therefore, as the water evaporates, the lakes become more salty and in some cases may dry up completely forming salt basins or playas especially in extremely arid environments. In Australia, for example, occasional rains may cause the dry salt basins to become lakes, and these then are called ephemeral salt lakes. Saline lakes are typically divided into two main types: athalassohaline and thalassohaline. The former refers to lakes which have a different chemical ratio of the main cations from what is found in seawater, while the latter refers to lakes with the cation ratio similar to seawater. Table 1 lists the sodium + potassium ion to calcium + magnesium ion ratios and summarizes the chemical composition of a variety of saline lakes thus showing the wide range of lake types possible. The data in Table 1 indicate that a subset of the athalassohaline lakes exists. These lakes have a high concentration of carbonate and therefore a high pH. These are the soda lakes or alkaline lakes. They, too, are found worldwide and are described below. No Antarctic lakes are listed in Table 1 because the surface waters are generally only very slightly saline (less than seawater) and it is only with depth, often greater than 10 m, that true saline waters are found. In general, it is the bottom waters that are hypersaline with little turnover to bring the saline waters to the surface. The chemical compositions of Antarctic lakes have been reviewed by Matsubaya et al. (1979). The reason for the differences in the chemical composition lies in the origins of the various types of lakes. The thalassohaline lakes were generally formed during the

Joachim Reitner & Volker Thiel (eds.), Encyclopedia of Geobiology, DOI 10.1007/978-1-4020-9212-1, # Springer Science+Business Media B.V. 2011

Utah, USA Israel 34.3 Iran 88 California, USA 29.5 Egypt 279.7 Bolivia 18.1 Saskatoon, Canada Gobi Desert Mongolia

Great Salt Lake – South Arm

Dead Sea Urmia Lake Mono Lake Wadi El Natrun Laguna Colorada

Muskiki

11

18.0

62.2

59

Calculation for this paper: (Na+ + K+)/(Ca++ + Mg++) Taylor et al. (1980) c Not listed d Not determined e Combined Na+ + K+

b

a

Seawater

Oygon

Australia Argentina Chile Utah, USA

Small lake near Koombekine Mar Chiquita Lake Laguna Amarga Great Salt Lake – North Arm

e

9.36 12.5 24 101

Location

Lake name

Na+

3.4

0.065 0.14 0.33 6.9

0.39

NL

8.0 1.1 1.5 NLc 1.85

K+

4.6

0.38 0.25 1.7 8.5

Mg++

95

15.7 16.7 8.2 175

Cl


0.42

0.02

NL

1.32

0.41

29

19.7

16.9

10 times its saturation) of Lake Van allows for in situ precipitation of aragonite, while in seawater, even though supersaturated, no spontaneous precipitation occurs. Rather, the supersaturation of seawater is kept low by the multitude of enzymatically biomineralizing organisms that are generally absent in soda lakes due to the high alkalinity.

Soda lake microbialites Those of the mature soda lakes that reach high supersaturation values (i.e., above SI 0.8, Kempe and Kazmierczak, 1990) produce cyanobacterial microbialites. Table 3 lists

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a few of them for which sufficient geochemical data are available to calculate saturation. The water analysis available for Andros Island originally do not show supersaturation, but the permineralizing cyanobacterial mats grow in ephemeral very shallow nonmarine inland lakes that apparently sustain growth only after an extensive evaporative phase. In case of one of the Niuafo‘ou caldera lakes (Vai Lahi), the current water composition also does not sustain in situ permineralization, and the stromatolites discovered there stopped growing about 200 years ago. Overall, soda lakes represent the best sites to study modern stromatolites, their structure, microbial ecosystem, and growth rates. Many of them have shown to produce structures strikingly similar to those of Precambrian microbialites.

Genesis of soda lakes These data presented here and in the cited papers show that soda chemistry can arise in various climatic, geologic, and morphologic settings. We find water bodies that are soda lakes in the making that will eventually acquire mature soda chemistry, such as Nemrut Crater Lake, a lake that originated only a few hundred years ago after the last eruption of the Nemrut caldera. Other crater lakes, such as Lake Taupo in New Zealand, will never reach this state, since they have an outlet. Astonishing is also the fact that Lake Kivu and Lake Tanganyika (both with an outlet) have soda lake characteristics (Soda Lake Index for Tanganyika: 1.5) and (at least in Lake Tanganyika) grow microbialites. It has long been stated that the type of chemistry is dependent on the proportions of the ions in the waters

Soda Lakes, Figure 2 Calcareous tufa towers in Mono Lake (top) and Pyramid Lake (bottom). These towers originally formed under water at inlets of groundwater.

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SODA LAKES

delivered to it (Garrels and Mackenzie, 1967). Eugster and Hardie (1978) constructed flow diagrams that show the various possibilities of brine evolution. In case of soda lakes, the criterion is the ratio of TA/(Mg + Ca). From the distributions of the soda lakes along the plate boundaries it appears that this ratio most likely arises in areas with fresh volcanic rocks. Here, biogenic CO2 and/or (see Lake Nemrut) volcanic CO2 can react with fresh silicates, mobilizing Na, K, Mg, and Ca, in the necessary proportions. Ongoing weathering within the crater lakes and their hydrothermal system (as in the case of Niuafo‘ou), or continuing evaporation in the terminal lakes will then lead to mature soda lake chemistry and to a CaCO3 supersaturation of around SI = 1 that will sustain microbialite growth. However, this is only a very broad picture. When looking at Satonda and Kauhako crater lakes, we find involvement of a second process that causes the alkalinity to rise. This is sulfate reduction. Both lakes are marinederived lakes, i.e., are filled originally with sulfate-rich seawater. Microbial disintegration of organic matter must replace the charge of the sulfate ion with that of bicarbonate ions according to C106 H263 O110 N16 P1 þ 53SO4 2
þ 14H2 O ! 53H2 S þ 106HCO3
þ HPO4 2
þ 16NH4 þ þ 14OH
This pathway of alkalinity production in anaerobic water bodies was coined “alkalinity pump” (Kempe, 1990; Kempe and Kazmierczak, 1994), supplementing the shift toward a soda chemistry, where sulfate is available.

Summary Soda lake waters show an excess of the total alkalinity over the charges of the alkaline earth ions magnesium and calcium ([HCO3
] + 2[CO32
] > 2[Mg2+] + 2[Ca2+]). They are characterized by high pH values (increasing with evaporation) and, eventually, precipitation of Na carbonates. Mature soda lakes that reach high supersaturation values with respect to calcite, aragonite, and dolomite (above SI 0.8) may produce cyanobacterial microbialites. The ubiquitous soda lakes in areas with fresh volcanic rocks and high weathering potential and the microbialites found therein gave rise to the Soda Ocean Hypothesis (see separate entry), stating that the early ocean, by following simple geological rules, should have been highly alkaline, possibly similar to present Lake Van. Bibliography Arp, G., Thiel, V., Reimer, A., Michaelis, W., and Reitner, J., 1999. Biofilm exopolymers control microbialite formation at thermal springs discharging into the alkaline Pyramid Lake, Nevada, USA. Sedimentary Geology, 126, 159–196. Benson, L. V., 1994. Carbonate deposition, Pyramid Lake subbasin, Nevada: 1. Sequence of formation and elevational distribution of carbonate deposits (tufas). Palaeogeography, Palaeoecology Palaeoclimatology, 109, 55–87.

Bischoff, J. L., Stine, S., Rosenbauer, R. J., Fitzpatrick, J. A., and Stafford, T. W., 1993. Ikaite precipitation by mixing of shoreline springs and lake water, Mono Lake, California, USA. Geochimica et Cosmochimica Acta, 57, 3855–3865. Donachie, S. P., Hou, S., Lee, K. S., Riley, C. W., Pikina, A., Kempe, S., Gregory, T. G., Bossuyt, A., Boerema, J., Liu, J., Freias, T. A., Malahoff, A., and Alam, M., 2004. The Hawaiian Archipelago: a microbial diversity hotspot. Microbial Ecology, 48, 509–520. Eugster, H. P., 1986. Lake Magadi, Kenya: a model for rift valley hydrochemistry and sedimentation? In Frostick, L. E. et al., (eds.), Sedimentation in the African Rifts. London: Geological Society, Special Publications 25, pp. 177–189. Eugster, H. P., and Hardie, L. A., 1978. Lake Magadi, Kenya: a model for rift valley hydrochemistry and sedimentation? In Lerman, A. (ed.), Physics and Chemistry of Lakes, New York: Springer, pp. 237–293. Garrels, R. M., and Mackenzie, F. T., 1967. Origin of the chemical composition of some springs and lakes. In equilibrium concepts of natural water systems. American Chemical Society Advances in Chemistry, 67, 222–242. Garret, D. E., 1992. Natural Soda Ash, Occurrences, Processing, and Use. New York: Van Nostrand Reinhold. Kazmierczak, J., and Kempe, S., 2006. Modern analogues of Precambrian stromatolites from caldera lakes of Niuafo‘ou Island, Tonga. Naturwissenschaften, 93, 119–126. Kazmierczak, J., Kempe, S., and Altermann, W., 2004. Microbial origin of Precambrian carbonates: lessons from modern analogues. In Eriksson, P., Altermann, W., Nelson, D., Mueller, W., and Catuneanu, O. (eds.), The Precambrian Earth: Tempos and Events. Amsterdam: Elsevier, pp. 545–564. Kempe, S., 1990. Alkalinity: the link between anaerobic basins and shallow water carbonates? Naturwissenschaften, 77, 426–427. Kempe, S., and Degens, E. T., 1985. An early soda ocean? Chemical Geology, 53, 95–108. Kempe, S., and Kazmierczak, J., 1990. Calcium carbonate supersaturation and the formation of in situ calcified stromatolites. In Ittekkot, V. A., Kempe, S., Michaelis, W., and Spitzy, A. (eds.), Facets of Modern Biogeochemistry, (Festschrift for E.T. Degens). Berlin: Springer-Verlag, pp. 255–278. Kempe, S., and Kazmierczak, J., 1993. Satonda Crater Lake, Indonesia: Hydrogeochemistry and biocarbonates. Facies, 28, 1–32. Kempe, S., and Kazmierczak, J., 1994. The role of alkalinity in the evolution of ocean chemistry, organization of living systems and biocalcification processes. In Doumenge, F. (ed.), Past and Present Biomineralization Processes. Considerations about the Carbonate Cycle. Bulletin de l’Institut océanographique, Monaco, no. spec. 13, 61–117. Kempe, S., and Kazmierczak, J., 1997. A terrestrial model for an alkaline martian hydrosphere. Planetary and Space Science, 45, 1493–1499. Kempe, S., and Kazmierczak, J., 2003. Modern soda lakes: model environments for an early alkaline ocean. In Müller, T., and Müller, H. (eds.), Modelling in Natural Sciences; Design, Validation and Case Studies. Berlin: Springer-Verlag, pp. 309–322. Kempe, S., Kazmierczak, J., Landmann, G., Konuk, T., Reimer, A., and Lipp, A., 1991. Largest known microbialites discovered in Lake Van, Turkey. Nature, 349, 605–608. Kraml, M., and Bull, A., 1998/1999. Sodaseen im Ostafrikanischen Graben – ihre Entstehung und Bedeutung. Berichte der Naturforschenden Gesellschaft zu Freiburg i. Br, 88/89, 85–118. Morse, J. W., Gledhill, W. K., and Millero, F. J., 2002. CaCO3 precipitation kinetics in waters from the Grand Bahama Bank: implications for the relationship between hydrochemistry and whitings. Abstracts of the 6th International Symposium on the Geochemistry of Earth’s Surface, May 20–24, 2002, Honolulu, Hawaii, pp. 138–140.

SODA OCEAN HYPOTHESIS

Parkhurst, D. L., Thorstenson, D. C., and Plummer, L. N., 1990. PHREEQE – A computer program for geochemical calculation. (Conversion and upgrade of the prime version of PHREEQE to IBM PC-compatible systems by Tirisanni, J. V., Glynn, P. D.,) US Geological Survey and Water Research Reports 80–96, 197 pp. Pegler, K., and Kempe, S., 1988. The carbonate system of the North Sea: determination of alkalinity and TCO2 and calculation of PCO2 and SIcal (Spring 1986). In Kempe, S., Liebezeit, G., Dethlefsen, V., and Harms, U. (eds.), Biogeochemistry and Distribution of Suspended Matter in the North Sea and Implications to Fisheries Biology. Mitteilungen aus dem GeologischPaläontologischen Institut der Universität Hamburg, SCOPE/ UNEP Sonderband, 65, pp. 35–87. Reimer, A., 1995. Hydrochemie und Geochemie der Sedimente und Porenwässer des hochalkalinen Van Sees in der Osttürkei, Faculty of Geosciences. Unpublished PhD Theses, University of Hamburg, 136 pp. Reimer, A., Landmann, G., and Kempe, S., 2009. Lake Van, Eastern Anatolia, hydrochemistry and history. Aquatic Geochemistry, 15, 195–222.

Cross-references Alkalinity Biofilms Calcite Precipitation, Microbially Induced Carbonates Cyanobacteria Divalent Earth Alkaline Cations in Seawater Dolomite, Microbial Microbial Mats Microbialites, Modern Soda Ocean Hypothesis Tufa, Freshwater

SODA OCEAN HYPOTHESIS Stephan Kempe1, Jozef Kazmierczak2 University of Technology, Darmstadt, Germany 2 Polish Academy of Sciences, Warszawa, Poland

1

Definition The “soda ocean hypothesis” (SOH) stands for the concept of an early (i.e., in essence Precambrian) alkaline or even highly “alkaline ocean,” in analogy to the chemistry of the present-day “soda lakes.” Soda ocean hypothesis (SOH) The SOH has been advanced in biology (e.g., Snyder and Fox, 1975) for biochemical reasons before it was developed in earth sciences for geochemical reasons (Kempe and Degens, 1985; Kempe et al., 1989; Kempe and Kazmierczak, 1994). In biology, the SOH rests on the observations that certain reactions considered essential for biogenesis would be favored by alkaline conditions (e.g., Abelson, 1966). One of those is the experimental observation that peptide bonds are more stable in alkaline than in acidic environments (e.g., Dose and Rauchfuss, 1972).

829

In earth sciences, the SOH rests on elemental mass balances, thermodynamic and kinetic arguments, and the analogy to modern soda lakes. These arguments are in short: 1. A CO2-rich atmosphere in the presence of water would react vigorously with the highly meta-stable, widespread, fine-grained glassy silicates from impacts and volcanic eruptions as would be present during the period of the terminal cataclysm in the Hadean. Carbonic acid weathering of silicates (“Urey Reaction”) will liberate cations in equivalence to the consumed carbonic acid, producing bicarbonate and carbonate as anions. 2. Continental silicate weathering today (aided by high soil PCO2 due to biological activity; PCO2 = CO2 Pressure) consumes annually 0.15 109t C (Hartmann et al., 2009); thus today’s volume of atmospheric CO2 would be consumed in about 5,000 years by the Urey Reaction. This rate suggests that a high PCO2 atmosphere is not stable on geological timescales. 3. Other primordial acids, such as HCl, have much less mass and therefore the primordial ocean must have had TA (total alkalinity [HCO3
] þ 2 [CO32
]) > [Cl
]. 4. Sulfur-based acids were not available in larger quantities since not enough free oxygen was in the system to allow formation of either sulfurous or sulfuric acids. 5. The entire mass of carbon available in crustal compartments (i.e., 65.5 1021 gC) as carbonates and organic carbon today could in principle have been dissolved as alkali carbonates in the oceans - as long as H2O and CO2 were degassed at similar rates - because this would amount to 48.5 gC/kg H2O while Na2CO3 is solvable at 53 gC/kg H2O (Kempe and Degens, 1985). 6. Observations of modern soda lakes show that their tributaries have inputs of [Naþ] þ [Kþ] > [Cl
] þ [SO42
] and [Ca2þ] þ [Mg2þ] < [TA] (Garrels and Mackenzie, 1967). When evaporated these waters will become alkaline because Ca and Mg carbonates (and sulfates in modern settings) will reach saturation first and Na and K carbonates will stay in solution causing increasingly higher alkalinity and pH. Soda lakes (i.e., lakes with an excess in Na-carbonates) almost exclusively occur today in or near volcanic regions. The early ocean (as well as possible primordial oceans on other planets such as Mars; Kempe and Kazmierczak, 1997) could have reacted the same way, storing CO2 as TA as quickly as CO2 was degassed from the mantle or deposited on Earth by comets. These arguments are based on simple acid–base reactions but have several derivatives: 1. The atmosphere could not have had a high enough PCO2 to counterbalance the lower insolation of the “faint early sun”. 2. In alkaline waters SiO2 can be kept in solution in high concentrations (at least in the primordial seas where

830

SODA OCEAN HYPOTHESIS

silica-sequestering organisms such as sponges, diatoms, silicoflagellates, and radiolarians were absent). 3. The higher the TA becomes, the lower the concentration of the free Ca-ion will become because of the fact that the IAP (the ion activity product) governing CaCO3 solubility, i.e., IAP = [Ca2þ]*[CO32
] is dominated by the carbonate ion. 4. In the absence of biologically controlled biomineralization (as was typical for the entire Precambrian until, at the onset of the Cambrian, enzymatic biocalcification evolved) the ocean must have been of a very high supersaturation, since the spontaneous precipitation of CaCO3-minerals is kinetically inhibited in natural settings. Saturation is most commonly expressed as the saturation index (SI), i.e., the quotient between the IAP and the solubility product of the mineral (KMineral). SI = log IAP/KMineral. A survey of nonenzymatic carbonate CaCO3-precipitating environments (Kempe and Kazmierczak, 1990; Kazmierczak and Kempe, 2006) shows that these environments have an SI of þ0.8 to þ1 (i.e., a 10-times supersaturation) while the modern ocean has a much lower supersaturation (North Sea SIcalcite average summer 1986 = 0.567  0.096; Pegler and Kempe, 1988) because it is controlled by enzymatic reactions and not by inorganic precipitation. Dolomite can be supersaturated to SI = 4, i.e., 10,000 times its solubility product in soda lakes (e.g., Kempe and Kazmierczak, 2007). This fact, i.e., the higher carbonate mineral supersaturation of the primordial ocean is today accepted widely (e.g., Grotzinger, 1990; Arp et al., 2001). Because the soda ocean has vanished and the present day halite ocean prevails, there must have been a transition between the two (Figure 1). This transition must have been driven by a process with a long time constant. This process

Soda Ocean Hypothesis, Figure 1 Simple representation of the timing of the rise and fall of the soda ocean due to the Urey reaction. The happy face marks the onset of multicellularity triggered by a massive increase of the free calcium ion towards the end of the Precambrian.

is thought to be the subduction of seawater with marine crust and sediments in subduction zones and the consecutive formation of continental crust. About 1 km3 of seawater is subducted per year today. Any compound dissolved in the ocean has therefore a half-life of about 1 billion years, depending on the rate of subduction. Thus the soda ocean could in fact have prevailed for several billion years before being gradually diminished. According to the soda ocean model (Kempe and Degens, 1985), Na and K would have been transferred into feldspars of continental granodiorites and the carbon would have been deposited as CaCO3 and kerogen on the continents where they are stored for longer times than deep sea deposits that are lost to subduction within less than 100 million years. The chloride subducted would be recycled as volcanic HCl, in a cycle quicker than continental weathering.

Consequences for biogenesis The alkaline model has also a series of consequence for biogenesis and the evolution of life in the first 3.5 billion years of Earth history. 1. Thermodynamic calculations show that in highly alkaline solutions (up to the saturation of Na2CO3), the free [Ca2þ] can be as low as 4*10–7 mol (Kazmierczak and Kempe, 2004), while, due to ion-pairs, total Ca is significantly higher. In the model calculation shown in Figure 2 seawater (i.e., a solution with high chloride and sulfate) was mixed with the natron-saturated solution based on the dilute water from Nemrut Crater Lake (
10 m), a “soda lake in the making” (see entry “Soda Lakes” Table 3) that leads to a free [Ca2þ] of 1.5*10–6 mol. Very low environmental Ca2þ concentrations are favorable for biogenesis because cells have to maintain a [Ca2þ] 10-fold increase in [Ca2þ] (Kempe and Kazmierczak, 1994). Measurements of Ca in fluid inclusions seem to give this facet of the SOH credibility (Brennan et al., 2004; Petrychenko et al., 2005).

Soda Ocean Hypothesis, Figure 2 PHREEQUE mixing experiment of modern seawater (left) with the natron saturated Nemrut Crater Lake water (see Table 2 in entry “Soda Lakes”) (right), illustrating the possible transition of an alkaline ocean (right) to one dominated by halite and gypsum (left). After linear mixing of constituents, the solutions are adjusted to have a calcite saturation index of +1 and a pPCO2 of 3.5 (roughly the present atmospheric PCO2). Note that the sharp drop in Ca concentration is accompanied by only a slight increase in alkalinity. Note also that the sulfate concentration would describe an exponential curve between 50 and 100% if subsamples were calculated for that section of the mixing experiment.

3.

4.

5.

6.

marine stromatolites are apparently growing predominantly by trapping and binding of allogene particles (e.g., Ginsburg, 1991). For three billion years cyanobacteria seem to be among the primary photosynthetic organisms. Most modern cyanobacteria are alkalophiles, suggesting that their evolution proceeded in alkaline environments. The development of eukaryotic organisms in the late Palaeoproterozoic may have also been triggered by the early ocean conditions: The gradual increase in the [Ca2þ] ambient concentration due to the slow transition between the soda and halite dominated oceans, could have caused the geochemical stress that might have influenced the evolution of a cell nucleus by protecting the genetic material against the disruptive Ca2þ-influx. And even the onset of multicellular organisms may have been triggered by the increasing [Ca2þ]. Experiments with sponges suggest that adhesion of cells can be triggered by an increase in [Ca2þ] in the environment (e.g., Kretsinger, 1977). Thus the appearance of faunas in Ediacara times may be the indication that the soda ocean was near its demise. The biomineralization that appears in many different taxa at the turn of the Precambrian/Cambrian is, in view of the SOH, a detoxification reaction to the

Counterarguments Some arguments against the SOH were brought forward. These, however, do not really disprove the SOH: 1. Models of early Earth often prescribe a long-lasting high PCO2-atmosphere to keep the oceans from freezing. However, it may also be argued that the lack of free oxygen in the oceans and atmosphere could have allowed for higher methane concentrations that prevented permanent freezing of Earth. 2. Möller and Bau (1993), Bau and Möller (1994) state that the water of Lake Van has a positive cerium anomaly but Precambrian carbonates (or BIFs) do not. However, these authors compare water with sediments. Measurements showed that Lake Van sediments do not exhibit the same anomaly as the water body (own data, unpublished). Outlook The acceptance of the SOH in the literature is mixed. Some authors agree with it in essence (Arp et al., 2001), others accept certain aspects (such as the high CaCO3supersaturation; Grotzinger and Kasting, 1993; Riding, 2000; or Mg/Ca ratios; Ries et al., 2008), and some doubt its relevance and follow own modeling lines (e.g., Morse and Mackenzie, 1998). Many papers deal with questions of the rising oxygen and sulfate, redox-reactions or certain sedimentary problems like BIF-formation or microbialite (stromatolite) structures without recurring to the overall chemical evolution of the ocean. Although the majority of researchers tend to ignore the SOH, it found entrance into several textbooks of general geology (Degens, 1989; Einsele, 1992; Bahlburg and Breitkreuz, 1998). Digital modeling appears to be not of much help - apart from stating thermodynamic conditions correctly - since the results depend on the adopted boundary conditions of the Hadean and Archean environment, which are as yet not well defined. For example, when prescribing certain PCO2 values to avoid freezing of oceans because of the faint early sun, then no highly alkaline ocean can evolve. Future research will have to show which one of the many models explains the past of our planet and its geological and biological evolution best. Summary Resting on elemental mass balances, thermodynamic and kinetic arguments, and the analogy to modern soda lakes, the soda ocean hypothesis (SOH) states that the Precambrian ocean was alkaline, with a total alkalinity >Ca þ Mg

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SODA OCEAN HYPOTHESIS

and a high supersaturation of carbonate minerals (SI ca. 1). The soda ocean was lost slowly by subduction of seawater and replaced by the present halite-dominated ocean toward the end of the Precambrian. This facilitated the development of multicellularity and biomineralization as detoxification reactions of life against the rising Caconcentrations. The SOH is also consistent with the widespread occurrence of stromatolites and cherts in the Precambrian.

Bibliography Abelson, P. H., 1966. Chemical events on the primitive Earth. Proceedings of the National Academy of Sciences USA, 55, 1365–1372. Altermann, W., Kazmierczak, J., Oren, A., and Wright, D. T., 2006. Cyanobacterial calcification and its rock-building potential during 3.5 billion years of earth history. Geobiology, 4, 147–166. Arp, G., Reimer, A., and Reitner, J., 2001. Photosynthesis-induced biofilm calcification and calcium concentrations in Phanerozoic oceans. Science, 292, 1701–1704. Bahlburg, G., and Breitkreuz, C., 1998. Grundlagen der Geologie. Stuttgart: F. Enke. Bau, M., and Möller, P., 1994. Präkambrische chemisch-sedimentäre Mineralisationen. Geowissenschaften, 12, 333–336. Brennan, S. T., Lowenstein, T. K., and Horita, J., 2004. Seawater chemistry and the advent of biocalcification. Geology, 32, 473–476. Carafoli, E., 1987. Intracellular calcium homeostasis. Annual Review of Biochemistry, 56, 395–433. Degens, E. T., 1989. Perspectives on Biogeochemistry. Berlin: Springer-Verlag. Dose, K., and Rauchfuss, H., 1972. On the electrophoretic behavior of thermal polymers of amino acids. In Rohlfing, D. L., Oparin, A. I. (eds.), Molecular Evolution: Prebiological and Biological. New York: Plenum Press, pp. 1–199. Einsele, G., 1992. Sedimentary Basins. Berlin: Springer-Verlag. Garrels, R. M., and Mackenzie, F. T., 1967. Origin of the chemical composition of some springs and lakes. In Equilibrium Concepts of Natural Water Systems. Advances in Chemistry, 67. American Chemical Society, pp. 222–242. Ginsburg, R. N., 1991. Controversies about stromatolites: Vices and virtues. In Müller, D. W., McKenzie, J. A., and Weissert, H. (eds.), Controversies in Modern Geology. London: Academic Press, pp. 25–36. Grotzinger, J. P., 1990. Geochemical model for Proterozoic stromatolite decline. In Knoll, A. H., and Ostrom, J. H. (eds.), Proterozoic Evolution and Environments, American Journal of Science (P. E. Cloud Special Volume), 290-A, pp. 80–104. Grotzinger, J. P., and Kasting, J. F., 1993. New constraints on Precambrian ocean composition. Journal of Geology, 101, 235–243. Hartmann, J., Kempe, S., Dürr, H. H., and Jansen, N., 2009. Global CO2-consumption by chemical weathering: What is the contribution of highly active weathering regions? Global and Planetary Change, 69, 185–194. Kazmierczak, J., and Kempe, S., 2004. Calcium build-up in the Precambrian sea: A major promoter in the evolution of eukaryotic life. In Seckbach, J. (ed.), Origins, Evolution and Biodiversity of Microbial Life. Dordrecht: Kluwer, pp. 329–345. Kazmierczak, J., and Kempe, S., 2006. Modern analogues of Precambrian stromatolites from caldera lakes of Niuafo‘ou Island, Tonga. Naturwissenschaften, 93, 119–126. Kempe, S., and Degens, E. T., 1985. An early soda ocean? Chemical Geology, 53, 95–108.

Kempe, S., and Kazmierczak, J., 1990. Calcium carbonate supersaturation and the formation of in situ calcified stromatolites. In Ittekkot, V. A., Kempe, S., Michaelis, W., and Spitzy, A. (eds.), Facets of Modern Biogeochemistry Festschrift for E.T. Degens on occasion of his 60th birthday, Berlin: Springer-Verlag, pp 255–278. Kempe, S., and Kazmierczak, J., 1994. The role of alkalinity in the evolution of ocean chemistry, organization of living systems and biocalcification processes. In Doumenge, F. (ed.), Past and Present Biomineralization Processes. Considerations about the Carbonate Cycle. Monaco: Bulletin de l’Institut océanographique, no. spec. 13, pp. 61–117. Kempe, S., and Kazmierczak, J., 1997. A terrestrial model for an alkaline martian hydrosphere. Planetary and Space Science, 45, 1493–1499. Kempe, S., and Kazmierczak, J., 2003. Modern soda lakes: Model environments for an early alkaline ocean. In Müller, T., and Müller, H. (eds.), Modelling in Natural Sciences; Design, Validation and Case Studies. Berlin: Springer-Verlag, pp. 309–322. Kempe, S., and Kazmierczak, J., 2007. Hydrochemical key to the genesis of calcareous non-laminated and laminated cyanobacterial microbialites. In Seckbach, J. (ed.), Algae and Cyanobacteria in Extreme Environments. Berlin: SpringerVerlag, pp. 241–264. Kempe, S., and Pegler, K., 1991. Sinks and sources of CO2 in coastal seas: the North Sea. Tellus, 43B, 224–235. Kempe, S., Kazmierczak, J., and Degens, E. T., 1989. The soda ocean concept and its bearing on biotic and crustal evolution. In Crick, R. E. (ed.), Origin, Evolution and Modern Aspects of Biomineralization in Plants and Animals Proceedings of the 5th International Symposium Biomineralization, Arlington, Texas, May, 1986, New York: Plenum Press, pp. 29–43. Kempe, S., Kazmierczak, J., Landmann, G., Konuk, T., Reimer, A., and Lipp, A., 1991. Largest known microbialites discovered in Lake Van, Turkey. Nature, 349, 605–608. Kretsinger, R. H., 1977. Evolution of the informational role of calcium in eukaryotes. In Wasserman, R. H., Corradino, R. A., Kretsinger, R. H., MacLennan, D. H., and Siegel, F. L. (eds.), Calcium Binding Proteins and Calcium Function. New York: North Holland Publishing, pp. 63–7. Kretsinger, R. H., 1983. A comparison of the roles of calcium in biomineralization and in cytosolic signaling. In Westbroek, P., and De Jong, E. W. (eds.), Biomineralization and Biological Metal Accumulation. Dordrecht: D. Reidel Publishing Co., pp. 123–131. López-Garcia, P., Kazmierczak, J., Benzerara, K., Kempe, S., Guyot, F., and Moreira, D., 2005. Bacterial diversity and carbonate precipitation in the microbialites of the highly alkaline Lake Van, Turkey. Extremophiles, 9, 263–274. Möller, P., and Bau, M., 1993. Rare-earth patterns with positive cerium anomaly in alkaline waters from Lake Van, Turkey. Earth and Planetary Science Letters, 117, 671–676. Morse, J. W., and Mackenzie, F. T., 1998. Hadean ocean carbonate geochemistry. Aquatic Geochemistry, 4, 301–319. Pegler, K., and Kempe, S., 1988. The carbonate system of the North Sea: Determination of alkalinity and TCO2 and calculation of PCO2 and SIcal (Spring 1986). In Kempe, S., Liebezeit, G., Dethlefsen, V., and Harms, U. (eds.), Biogeochemistry and Distribution of Suspended Matter in the North Sea and Implications to Fisheries Biology, Mitteilungen aus dem GeologischPaläontologischen Institut der Universität Hamburg, SCOPE/ UNEP Sonderband, 65, pp. 35–87. Petrychenko, O. Y., Peryt, T. M., and Chechel, E. I., 2005. Early Cambrian water chemistry from fluid inclusions in halite from Siberian evaporates. Chemical Geology, 219, 149–161.

SOILS

Riding, R., 2000. Microbial carbonates: the geological record of calcified bacterial-algal mats and biofilms. Sedimentology, 47, 179–214. Ries, J. B., Anderson, M. A., and Hill, R. T., 2008. Seawater Mg/Ca controls polymorph mineralogy of microbial CaCO3: A potential for calcite-aragonite seas in Precambrian time. Geobiology, 6, 106–119. Snyder, W. D., and Fox, S. W., 1975. A model for the origin of stable protocells in a primitive alkaline ocean. Biosystems, 7, 222–229.

Cross-references Alkalinity Biofilms Calcite Precipitation, Microbially Induced Carbonates Cyanobacteria Divalent Earth Alkaline Cations in Seawater Dolomite, Microbial Microbial Mats Microbialites, Modern Soda Lakes Tufa, Freshwater

SOILS Erika Kothe Friedrich Schiller University Jena, Jena, Germany

Synonyms Pedosphere Definition Soil: refers to the product of mineral weathering and secondary mineral formation, (microbial) mineralization, humus formation, and the resulting element mobilization/immobilization in the upper Earth crust in a pedogenetic process involving chemical, physical, and biological activities. Introduction Soil is the basis of terrestrial life, particularly for agriculture, forestry, and generally land-use by man (Driessen et al., 2001). It is also the largest terrestrial ecosystem dominated by high numbers of microorganisms and soilliving animals as well as the root systems of plants. The biota within this ecosystem, not only interact but also actively shape their environment (Fiedler et al., 2002). This can be easily seen with root systems or earthworm/ mole tunnels, but the metabolic activities of microorganisms by far exceed these visible alterations. Microbes are essential for the decomposition of plant litter followed by humification. The pedogenesis differs with respect to base rock material, climatic conditions, and other factors leading to soil series of different soil types. Root exudates as well as microbiological acidification and release of

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chelating agents take part in mobilization and immobilization processes of mineral elements, essential or ecotoxically relevant alike, and thus are major players in import into food chains, and input into water paths or as volatiles into the atmosphere.

Soil function The soils of the world can be characterized from a more structural or a more functional point of view (FAOUNESCO, 2002), and the geobiologically important functions involving metabolically active soil dwellers are characterized with reference to microbiology, botany and agriculture, and zoology. Soil function is essential for plant nutrition. The capacity to solubilize nutrients like P and minerals including K, Na, Mg, Ca, and trace elements, as well as availability of S or N are largely the product of microbial mobilization from minerals, microbial degradation of organic matter, or fixation from air (e.g., for nitrogen). Soil mineralogy largely determines the buffering capacity for minerals adsorbed to different fractions (Jones and Bassington, 1998). The sustainability of microbial activity is connected to water availability for growth, and water storage and availability are also major functions for plant growth. Detrimental processes of soil degeneration like soil acidification, desertification, clay mineral destruction, or decrease of organic contents are global problems mankind experiences which will have a large impact on socioeconomic scales. Agriculture and nutrition in modern systems are coupled to the use of fertilizers and plant protection chemicals which can constitute a threat to the water pathways, if management of agricultural use is not performed according to the laws and provisions on soil and water protection, good agricultural practice, or integrated farming, and monitoring has to be included to ensure best practice. Soil density increases due to the use of heavy machinery which leads to soil degradation in high-yield agricultural systems the world over. In order to assess possible problems with land-use and soil degeneration, soil science has provided a very good reference system to classify and monitor soils based on soil mechanics, soil physics, soil genesis, soil composition, and soil functions reviewed in different contexts below (IUSS Working Group WRB, 2006). As soil can be regarded as a major sink for carbon depending on land-use, this might become more important as climate change discussions are gaining impact. Generally, soils and larger views on landscape should be regarded as eminent players in global cycles (Meir et al., 2006). Soil texture The soil texture can be divided in the proportion of sand (S: 63 μm–2 mm), silt (U: 2–63 μm), and clay (T: smaller than 2 μm) after removing skeleton particles (larger than 2 mm). While sand soils have more than 70 volume % of sand, clay soil is mainly composed of clay (>50%) with

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only small portions of sand (10/mm);

SULFATE-REDUCING BACTERIA

(e) association of SFF with or occurrence within SFF of other morphological bioforms such as fossil Gallionella or Metallogenium-like structures; (f) presence of vertically oriented elements (may resemble stalactites) resulting from mineral encrustation of filaments, filament strands, or mats while they are still flexible; and (g) inferred environmental conditions compatible with microbial growth, such as an abundant energy source and temperatures between ambient and 50 C) in wellcrystallized veins or masses and at low temperatures (50% of the material with a particle size 80% of the thin sections) followed by mollusc and brachiopods shells, fragments of trilobites, and hyalosteliid spicules (Figure 5b). However, the abundance of skeletal components was low in both settings, although they were volumetrically more important and diverse on the mud mounds. Most of the variations could be related with taphonomic bias and differences in available habitats (cryptic spaces, soft, firm, and hard substrates) more suitable in/on mud mounds. The systematic analysis of the biotic content from the off mound record was developed by Jeffery and Stanton (1996), and four component assemblages were established from deep to more shallow conditions:
Assemblage I: crinoid/echinoderm debris, fenestellid

hash, and ostracods

WAULSORTIAN MUD MOUNDS

C

B

Aphotic

Waulsortian Phases

D

Photic

Fe n C este r in t r Fe oid ate ne s sh ee O str st a t r t Br aco e h ac d as O h/b s h th iv e Si r b alve m ry s Tr ple ozo ilo fo a Ec bite ram ns h s s M in o o r id Fi ava spi la m n e m H e n min s ya t id s s Se lost r p e li i Pe uli d lo d C ids tube ya g s G nop eop as y e t t Ao trop es al uj od Pl ga s u r liid In iloc s tra ul C cla ar f ry st or p M tal s am s ic g a r it l M is co ic e d a r M itis ce ting ic e d m s r C itis wa ent lo e d ll ro s ph gra yt in es s Light Depth Energy

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A

Waulsortian Mud Mounds, Figure 4 Qualitative distribution of components through the Waulsortian Phases (A, B, C, and D). Redrawn and modified from Lees and Miller (1995).
Assemblage II: I þ stacheiins (possible algae), rare

salebrids (possible bryozoan), and commonly abundant sponge spicules
Assemblage III: II þ plurilocular foraminifera
Assemblage IV: III þ green algae

Biologically controlled contribution The dominant macrobiota in the Waulsortian mud mound was formed by heterozoan assemblages, which never constituted a skeletal framework. Photosynthetic groups such as calcareous algae were very rare and uncommon to colonize these mud mounds. Crinoids and bryozoans have played an important role when they colonized and stabilized the slopes and mound surfaces. As skeletal producers, they formed favorable substrates (encrusters, cement nucleation, etc.) and they had also relative importance as allomicrite bafflers. Thus, the endemic macrofauna was composed by crinoids, fenestrate bryozoans, and sponges (Figure 6). The metabolism and decay of siliceous sponges could also favored the bacterial activity producing organomicrites, which would be added to the main autochthonous carbonate production in these mud mounds (biologically induced mechanism are argued by most authors). Opportunistic epi- and infauna biota could colonize the locally soft mound substrates and after their death contribute to the biodegradation with the skeletal debris (Figure 6). Carbonate muds The most important component in the Waulsortian mud mounds is the carbonate mud, paradoxically during many

decades its nature has been not considered or specifically studied. In fact, the origin of the carbonate muds and its ability to maintain steep slopes has been subject of much debate during years. Some authors have advocated for an external source followed by some kind of posterior accumulation mechanisms (hydrodynamic, baffling by bryozoans and crinoids, trapping and binding by bryozoans or other organisms). From the 1980s most part of the authors suggest a probable microbial origin for the carbonate mud, and a dominant in situ carbonate mud production is assumed. The carbonate muds appear forming mudstone to packstone, although most works coincide in wackestone as the commonest texture. They are highly organized displaying successive generations of carbonate muds in geopetal relationship. They have been separated in primary muds (M1) and later muds (M2, M3, M4, etc.), all forming together the known polymuds (name coined by Lees and Miller, 1985, after the “multicomponent mudstones” concept of Lees (1964)). The earliest mud generation (M1) can represent less than 50% of carbonate mud volume. Lees and Miller (1995) subdivided the primary muds into three different subtypes (Figure 6):
(M1a) optically dense micrites, often with filaments
(M1b) paler biomicrites
(M1c) peloidal micrites

A biofilm model has been proposed for the in situ primary mud production (Figure 6). The biofilm formed by bacterial cells, cyanobacterial filaments, and bacterial extracellular polymeric substances (EPS), induced the precipitation of high-magnesium calcite micrites (dense

WAULSORTIAN MUD MOUNDS

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Waulsortian Mud Mounds, Figure 5 (a) Muleshoe mud mound, Sacramento Mountains, New Mexico. Mud mound growth phases following Kirkby and Hunt (1996). (b) Components distribution of 26 elements from both off mound levels and mud mounds (Alamogordo Member mounds and Muleshoe mound – Alamogordo, Nunn, and Tierra Blanca Members). Modified from Ahr and Stanton (1996). % Frequency of occurrence; R = rank position.

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WAULSORTIAN MUD MOUNDS

Waulsortian Mud Mounds, Figure 6 Biofilm model with the process-response mechanisms proposed to Waulsortian mud mounds. (Simplified and modified after Lees and Miller, 1995.)

WAULSORTIAN MUD MOUNDS

and peloidal ones). Biomicrites were formed by both reworking of dense or/and peloidal micrites and the skeletal debris input (Figure 6). Additionally, the biofilm could be used by other mud mound colonizers as a source of organic carbon. Carbonate mud production was not only reduced to the active biofilm mound surface, but also continued below the surface favored by the biofilm degradation during early burial (Figure 6). Dewatering, shrinkage, carbonate dissolution, and reworking processes produced the collapse of carbonate mud matrix. The collapse and successive reorganizations of the carbonate muds through the cavity system resulted in the final polymud fabrics. Later fractures and fissures could also have favored sediment inputs through the mound void system (Figure 6). There are no available data on the quantitative distribution/contribution of the different primary muds through the Waulsortian phases. Filaments, characteristic in dense micrites (M1a) donot occur in phase A, whereas they are abundant in the rest of phases. As well as, cavities from phase D contain important volumes of geopetal peloids intercalated with early calcite cements. Thus an early, deep, nonfilamentous, non-photosynthetic cyanobacterial community has been suggested for the carbonate mud production in the Waulsortian phase A (Miller, 1986). In the same way, several microbial communities may have controlled the distribution/production of the different primary non-reworked mud textures (dense or peloidal micrites). The distribution and typologies of carbonate mud, from the transitional facies to Waulsortian mud mound nucleation, were analyzed by Devusyst and Lees (2001), in western Ireland. Precursor muds are differentiated in function by their grumous or non-grumous character, bioclastic content, and contact relationship. They were grouped into two major types with six subtypes:
Grumous types on basis of their apparent optical den-

sity, with three intergrading subtypes (codified as 1, 2 and 3). Types 1 and 2 are normally poor in skeletal debris; however, the dark optically dense type 3 contains abundant sponge spicules. All of them have been interpreted as automicrites. The grumous types are equivalent to the peloidal micrites (M1c) of Lees and Miller (1985, 1995).
Non-grumous types: (1) gradational (into grumous muds) bioclastic wackestones; (2) distinct bioclastic wackestones; and (3) packstone matrix. The first subtype is also interpreted as automicrite whereas the others are seen as loose sediment. The different precursor muds and other features were analyzed by correspondence analysis and a compositional gradient or trend was detected (Figure 7). The precursor muds display different textural and genetic types and culminate in the Waulsortian polymuds that contain both grumous and non-grumous muds (mainly non-grumous).

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Waulsortian Mud Mounds, Figure 7 Observed relay pattern resulting from correspondence analysis of the precursor muds, polymuds, and associated features. Simplified and modified from Devusyst and Lees (2001). Vertical scales of boxes ranges from 0 up to 1, thus grey areas represent the frequency of presence. CFC = crypto-fibrous calcite; w = wackestone.

Open space structures Waulsortian mud mounds were originally interpreted as reefs by Dupont (1881) who coined the term Stromatactis in 1881 to describe and interpret the “sparry masses” from Devonian mud mounds as recrystallized fossils. The stromatactis are a special type of cavities whose origin is still controversial. They have a characteristic shape, with a flat to undulose floor and irregular digitate roof and are filled by centripetal calcite cement, although internal sediment can also occur. Other terms like stromatactoid or stromatactis-like cavity are commonly found in the literature to describe similar features. They are common through the Precambrian up to the Jurassic mud mounds, (Bosence and Bridges, 1995) although can also appear in other types of limestones not just in muddy ones. A different hypothesis has been proposed to explain the stromatactis, which can be grouped in organic or inorganic models (see reviews by Bathurst, 1982; Flajs and Hüssner, 1993; Monty, 1995; Flügel, 2004, p. 194). In Belgium, some mud mounds facies contain those described as undulating or tortuous “vein bleues” which were used, for many years, as a traditional criterion to characterize the Waulsortian facies and but was later abandoned (Lees, 1988). Sheet-form cavities, similar as known zebra cavities, just few centimeters thick but several meters in length, are also typical in Waulsortian mud mounds. Their roofs

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WAULSORTIAN MUD MOUNDS

are not clearly associated with skeletal support and they have been interpreted as the result of dewatering and shear stress processes in the firm, gel-like consistence muds. Stromatactis, the sheet-form and irregular shelter cavities have been considered early-formed cavity systems (Lees and Miller, 1995). Dissolution as well as physical processes like fracturing and internal erosion produce secondary cavities included fissures (Figure 6). The size of the cavities tends to decrease through B and C phases and gets much complex in the D than in the others. Thus, “sparry” fabrics are common in phase A, whereas in B and C they tend to be muddier. The shape and heterogeneity of the cavities seem to mainly be related with the skeletal content as well as the different carbonate mud textures and their distribution. Fillings as polymuds, cements (marine fibrous calcites are common), and their diagenetic sequences are also diverse (Meyers, 1974; Miller, 1986).

Conclusions The understanding of the Waulsortian reefs, carbonate mudbanks, mounds, buildups, or mud mounds has evolved from the last two centuries. The application of Waulsortian term is generally related to a group of diagnosis criteria: complex carbonate mud fabrics (polymuds), stratigraphic position (Middle-Upper Tournaisian-Lower Viséan), facies, component assemblages (Waulsortian phases), and the associated macrofaunas. The Mississippian seas were colonized by a wide bioconstruction spectrum and not by Waulsortian mud mounds alone. Bacteria and cyanobacteria are suggested as primary producers in Waulsortian microbial mud mounds. The hypothesis about the distribution of Waulsortian mud mounds has been related with upwelling areas and methane seeps. Carbon sources and convincing models to explain such large volumes of radiaxial fibrous calcite cements and continuous regional production of polymuds are still unrevealing. Most part of the studies has been focussed on the allochems distribution that has been considered in detail by many authors. Few works are specifically dedicated to the analysis of the carbonated mud, main component in the Waulsortian mud mounds. However, more work would be necessary to analyze, quantify, and decipher the relationship between the allomicrite input and the automicrite production in mud mounds, particularly in Waulsortian ones, where clear microbialite frameworks are not always evident. Summary Waulsortian mud mounds are probably the most famous carboniferous bioconstructions. They were described first from Belgium outcrops, in the locality of Waulsort, two centuries ago. Later, they have been recognized and analyzed from several areas of Europe and North America.

The growth of these bioconstructions was controlled by the dominant autochthonous production of carbonate mud which has been explained by the activity of marine microbial benthic communities. The associated biota which colonized mud mound substrates was composed by heterozoan assemblages dominated by fenestrate bryozoans and crinoids. Detailed studies of these component assemblages have been a key to reconstruct regional gradients from deep subtidal, aphotic marine environments through the photic zone. Today, there are no equivalent analogs to compare with such a spectacular mud mound development during the Tournaisian-lowermost Viséan period. Thus, they represent an excellent record to reconstruct and understand the paleoenvironmental and paleoecological relationships between microbial and nonmicrobial benthic communities through deep ramp to basin settings along the northern hemisphere during Lower Mississippian times.

Bibliography Ahr, W. M., and Stanton, R. J. Jr., 1996. Constituent composition of early Mississippian carbonate buildups and their level-bottom equivalents, Sacramento Mountains, New Mexico. In Strogen, P., Somerville, I. D., and Jones, G. Ll. (eds.), Recent Advances in Lower Carboniferous Geology. London: Geological Society Special Publication, Vol. 107, pp. 83–95. Aretz, M., and Webb, G. E., 2006. Western European and eastern Australian Mississippian shallow-water reefs: a comparison. In Wong, Th. E., (ed.), Proceedings of the XVth International Congress on Carboniferous and Permian Stratigraphy. Utrecht, the Netherlands: Royal Netherlands Academy of Arts and Sciences, pp. 433–441. Bathurst, R. G. C., 1982. Genesis of stromatactis cavities between submarine crusts in Paleozoic carbonate mud mounds. Journal Geological Society of London, 139, 165–181. Bosence, D. W. J., and Bridges, P. H., 1995. A review of the origin and evolution of carbonate mud-mounds. In Monty, C. L. V., Bosence, D. W. J., Bridges, P. H., and Pratt B. R. (eds.), Carbonate Mud-mounds: Their Origin and Evolution. Oxford: Blackwell Science for the International Association of Sedimentologists, Special Publication, 23, pp. 3–9. Bridges, P. H., Gutteridge, P., and Pickard, N. A. H., 1995. The environmental setting of early carboniferous mud-mounds. In Monty, C. L. V., Bosence, D. W. J., Bridges, P. H., and Pratt, B. R. (eds.), Carbonate Mud Mounds their Origin and Evolution. Special Publication International Association of Sedimentologists, 23, pp. 171–190. Devusyst, F.-X., and Lees, A., 2001. The initiation of Waulsortian buildups in Western Ireland. Sedimentology, 48, 1121–1148. Dupont, E., 1863. Sur le Calcaire Carbonifère de la Belgique et du Hainaut franBais. Bulletin de l’Académie Royale des Sciences des Lettres et des Beaux-arts de Belgique, 2 série, 15, 86–137. Dupont, E., 1881. Sur l’origine des calcaires de la Belgique. Académie Royale des Sciences de Belgique, 3 serie, 5, 264–280. Flajs, G., and Hüssner, H., 1993. A microbial model for the Lower Devonian stromatactis mud mounds of the Montagne Noire (France). Facies, 29, 179–194. Flügel, E., 2004. Microfacies of Carbonate Rocks, Analysis, Interpretation and Application. Berlin: Springer, pp. 976. Jeffery, D. Ll., and Stanton, R. J. Jr., 1996. Biotic gradients on a homoclinal ramp: the Alamogordo member of the Lake Valley

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formation, Lower Mississippian, New Mexico, USA. In Strogen, P., Somerville, I. D., and Jones, G. Ll. (eds.), Recent Advances in Lower Carboniferous Geology. Geological Society Special Publication, 107, pp. 111–126. Kirkby, K. C., and Hunt, D., 1996. Episodic growth of a Waulsortian buildup: the Lower Carboniferous Muleshoe Mound, Sacramento Mountains, New Mexico, USA. In Strogen, P., Somerville, I. D., and Jones, G. Ll. (eds.), Recent Advances in the Lower Carboniferous Geology. Geological Society of London Special Publication, 107, pp. 97–110. Lees, A., 1964. The structure and origin of the Waulsortian (Lower Carboniferous) “reefs” of west-central Eire. Philosophical Transactions of the Royal Society of London, Series B, 247, 483–531. Lees, A., 1988. Waulsortian reefs: the history of a concept. Mémoires de l’Institut Géologique de l’Université de Louvain, 34, 43–55. Lees, A., Hallet, V., and Hibo, D., 1985. Facies variation in Waulsortian buildups. Part 1. A model from Belgium. Geological Journal, 20, 133–158. Lees, A., and Miller, J., 1985. Facies variation in Waulsortian buildups. Part 2. Mid-Dinantian buildups from Europe and North America. Geological Journal, 20, 159–180. Lees, A., and Miller, J., 1995. Waulsortian banks. In Monty, C. L. V., Bosence, D. W. J., Bridges, P. H., and Pratt, B. R. (eds.), Carbonate Mud-mounds, their Origin and Evolution. International Association of Sedimentologists Special Publication, 23, pp. 191–271. Meyers, W. J., 1974. Carbonate cement stratigraphy of the Lake Valley Formation (Mississippian), Sacramento Mountains, New Mexico. Journal of Sedimentary Petrology, 44, 837–861. Miller, J., 1986. Facies relationships and diagenesis in Waulsortian mud mounds from the Lower Carboniferous of Ireland and N England. In Schroeder, J. H., and Purser, B. H. (eds.), Reef Diagenesis. Berlin: Springer, pp. 311–335. Monty, C. L. V., 1995. The rise and nature of carbonate mudmounds: an introductory actualistic approach. In Monty, C. L. V., Bosence, D. W. J., Bridges, P. H., and Pratt, B. R. (eds.), Carbonate Mud Mounds, their Origin and Evolution. International Association of Sedimentologists Special Publication, 23, pp. 11–48.

Cross-references Bacteria Biofilms Calcite Precipitation, Microbially Induced Cyanobacteria Extracellular Polymeric Substances (EPS) Microbial Communities, Structure, and Function Microbialites Mud Mounds Reefs

WHALE AND WOOD FALLS Steffen Kiel University of Göttingen, Göttingen, Germany

Synonyms Biogenic substrates; Food falls; Nekton falls; Organic falls; Sunken wood

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Definition Whale fall: Carcass of a large marine mammal of the order Cetacea (whales) that has sunken to the seafloor. Wood fall: A piece of wood that has sunken to the seafloor. History The first examples of specialized invertebrates inhabiting bones and wood in the deep sea (qv) were recognized during the deep-sea expeditions in the early to mid 1900s. Because these samples were recovered by dredging, the complexity of whale- and wood-fall ecosystems was not recognized before the advent of deep-sea research vessels. Pioneering experimental work on wood-falls was carried out in the early 1970s by Ruth Turner (1914–2000), using the deep-sea research vessel Alvin. This research came to a halt in the late 1970s when deep-sea hydrothermal vent systems (qv) and their fascinating fauna were discovered and Alvin was occupied by exploring these systems. The first deep-sea whale-fall ecosystem was accidentally discovered by Craig Smith during an Alvin dive in 1987 and consisted of a whale skeleton associated with many chemosymbiotic taxa that were also known from hydrothermal vents (Smith et al., 1989). Nowadays, whale-fall ecosystems are mainly investigated by sinking stranded whales which are then studied by submersibles and/or ROVs. The last major discovery in this field was Osedax, a “bone-eating” siboglinid worm living in symbiosis with bacteria that degrade bone lipids and cartilage (Rouse et al., 2004). Bivalve-bored wood has been known from the fossil record for centuries, but reports on other invertebrate groups that utilized wood falls in the geologic past are rare, and entire fossil wood-fall ecosystems have only been described very recently (Kiel and Goedert, 2006b; Kiel et al., 2009). Fossil examples of deep-sea whale-fall ecosystems were discovered soon after the discovery of the modern ones and are especially common in uplifted deep-water sediments around the North Pacific margin (Squires et al., 1991; Kiel and Goedert, 2006a). Similar ecosystems were recently found on two late Cretaceous plesiosaur (a marine reptile) skeletons in Japan (Kaim et al., 2008). Whale falls Whale falls are the basis for the most species-rich deep-sea ecosystem, with more than 400 species known to date (Baco and Smith, 2003). Whale falls pass through three ecologic stages that vary in length and can overlap. At first the flesh of the whale carcass is removed by sleeper sharks, hagfish, and other scavenging deep-sea fishes. In the second stage small annelids and crustaceans consume the remaining particulate organic matter. The main source of nutrients during the third stage is the hydrogen sulfide (H2S) generated by the anaerobic decay of bone lipids, which is utilized by species harboring chemotrophic endosymbionts or by species that graze on sulfur-oxidizing

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bacteria (Smith and Baco, 2003). While this “sulfophillic stage” can last up to 100 years in oxygen-poor ocean basins, it can be as short as a few years in areas where Osedax rapidly consumes and destroys the bones (Braby et al., 2007; Fujiwara et al., 2007). Whale bones consist of up to 65 wt% of lipids. These lipids are consumed by sulfate-reducing bacteria (qv) at the oxic–anoxic interface in the bone, using seawater as sulfate source and emitting H2S as metabolic byproduct. This H2S is then consumed by Beggiatoa (qv) and other sulfur-oxidizing bacteria on the surface of the skeleton (Figures 1 and 2). Bone lipids and other whale remains entering the, or covered by, sediment are also consumed in this way. The resulting H2S is either consumed by bacterial mats surrounding the whale carcass, or by invertebrates harboring sulfur oxidizing bacteria. These invertebrates include siboglinid tube worms, solemyid,

Whale and Wood Falls, Figure 1 Whale skeleton in the deep sea, covered by a large bacterial mat; photo by Craig R. Smith.

Sulfuroxidizing bacteria

SO42–

Bivalves with sulfur-oxidizing endosymbionts

H2S

Lipids SO42–

Seawater sediment

H2S

Bone

Lipids

Sulphate-reducing bacteria

Whale and Wood Falls, Figure 2 Generalized geobiologic processes at whale falls. (Modified from Smith and Baco, 2003.)

lucinid, and vesicomyid clams, and bathymodiolin mussels. Bacteria-grazing taxa include provannid, limpet-like and various other small-sized gastropods (“skeneimorphs”), and polynoid annelids (Smith and Baco, 2003). Fossil whale fall communities are known from to Pliocene sediments. They preserve mollusk associations resembling those at modern whale falls, and fossil traces of Osedax (Kiel et al., 2010).

Wood falls Wood falls in the deep sea support complex ecosystem of 40 or more invertebrate species, some of which are endemic to this habitat. The wood is utilized by several xylophagous (wood-eating) invertebrate groups. Most important are wood-boring bivalves of the heterodont families Teredinidae (also known as shipworms; mainly in shallow water) and Xylophagainae (mainly in deep water below 100 m) which rapidly consume and destroy the wood, presumably aided by cellulolytic symbiontic bacteria (Turner, 1973; Distel and Roberts, 1997). Shallow water wood falls are also utilized by wood-boring isopods of the family Limnoriidae. In deeper water exists a small number of xylophagous gastropod limpets and chitons (polyplacophorans), which are preyed upon by gastropods, annelids, and crustaceans. The microbial fauna on wood falls consists of a variety of fungi and bacteria (Kohlmeyer and Kohlmeyer, 1979; Palacios et al., 2006) and these microbes are grazed upon by gastropods and chitons. Chemosymbiotic species which are phylogenetically related to those at whale falls are also found on sunken wood. They probably rely on sulfide from the anaerobic decay of fecal pellets of the shipworms, or from the decaying wood itself (Figures 3 and 4; Kiel and Goedert, 2006b; Pailleret et al., 2007). The fossil record of deep-sea wood-fall communities extends back into the late Cretaceous. Summary Whale and wood falls in the deep sea harbor species-rich ecosystems that rely to a certain extent on H2S from the decay of the bones and the wood. Many of the species inhabiting these habitats are endemic. Especially the chemosymbiotic and bacteria-grazing taxa are phylogenetically related to those living at hydrothermal vents (qv) and cold seeps (qv). The main H2S source for the chemosymbiotic taxa at whale falls is the anaerobic breakdown of bone lipids by sulfate-reducing bacteria. In the case of the wood falls, it is the anaerobic decay of excrements of wood-boring bivalves (Xylophagainae). Wood-fall communities with a fauna that is taxonomically comparable to the modern wood-fall fauna are known from the late Cretaceous. Whale-fall communities are as old as ocean-going whales (late Eocene). Similar ecosystems with bacteria-grazing gastropods on plesiosaur bones are known from the late Cretaceous.

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Whale and Wood Falls, Figure 3 Thin section image of a fossil wood fall. White arrows indicate fecal pellets of wood-boring xylophagain bivalves, black arrow indicates borehole filled by authigenic carbonate.

Whale and Wood Falls, Figure 4 Generalized geobiologic processes at wood falls. (Redrawn from Kiel, 2008.)

Bibliography Baco, A. R., and Smith, C. R., 2003. High species richness in deepsea chemoautotrophic whale skeleton communities. Marine Ecology Progress Series, 260, 109–114. Braby, C. E., Rouse, G. W., Johnson, S. B., Jones, W. J., and Vrijenhoek, R. C., 2007. Bathymetric and temporal variation among Osedax boneworms and associated megafauna on whale-falls in Monterey Bay, California. Deep-sea Research, I, 54, 1773–1791. Distel, D. L., and Roberts, S. J., 1997. Bacterial endosymbionts in the gills of the deep-sea wood-boring bivalves Xylophaga atlantica and Xylophaga washingtona. Biological Bulletin, 192, 253–261. Fujiwara, Y., Kawato, M., Yamamoto, T., Yamanaka, T., Sato-Okoshi, W., Noda, C., Tsuchida, S., Komai, T., Cubelio, S. S., Sasaki, T., Jacobsen, K., Kubokawa, K., Fujikura, K., Maruyama, T., Furushima, Y., Okoshi, K., Miyake, H.,

Miyazaki, M., Nogi, Y., Yatabe, A., and Okutani, T., 2007. Three-year investigations into sperm whale-fall ecosystems in Japan. Marine Ecology, 28, 219–232. Kaim, A., Kobayashi, Y., Echizenya, H., Jenkins, R. G., and Tanabe, K., 2008. Chemosynthesis-based associations on Cretaceous plesiosaurid carcasses. Acta Palaeontologica Polonica, 53, 97–104. Kiel, S., 2008. Fossil evidence for micro- and macrofaunal utilization of large nekton-falls: examples from early Cenozoic deepwater sediments in Washington State, USA. Palaeogeography, Palaeoclimatology, Palaeoecology, 267, 161–174. Kiel, S., and Goedert, J. L., 2006a. Deep-sea food bonanzas: early Cenozoic whale-fall communities resemble wood-fall rather than seep communities. Proceedings of the Royal Society B, 273, 2625–2631. Kiel, S., and Goedert, J. L., 2006b. A wood-fall association from Late Eocene deep-water sediments of Washington State, USA. Palaios, 21, 548–556. Kiel, S., Amano, K., Hikida, Y., and Jenkins, R. G., 2009. Wood-fall associations from Late Cretaceous deep-water sediments of Hokkaido, Japan. Lethaia, 42, 74–82. Kiel, S., Goedert, J. L., Kahl, W. -A., and Rouse, G. W., 2010. Fossil traces of the bone-eating worm Osedax in early Oligocene whale bones. Proceedings of the National Academy of Sciences of the USA, 107, 8656–8659. Kohlmeyer, J., and Kohlmeyer, E., 1979. Marine Mycology: The Higher Fungi. New York: Academic Press. Pailleret, M., Haga, T., Petit, P., Privé-Gill, C., Saedlou, N., Gaill, F., and Zbinden, M., 2007. Sunken wood from the Vanuatu Islands: identification of wood substrates and preliminary description of associated fauna. Marine Ecology, 28, 233–241. Palacios, C., Zbinden, M., Baco, A. R., Treude, T., Smith, C. R., Gaill, F., Lebaron, P., and Boetius, A., 2006. Microbial ecology of deep-sea sunken woods: achieving quantitative measurements for bacterial biomass and cellulolytic activities. Cahiers de Biologie Marine, 47, 415–420.

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Rouse, G. W., Goffredi, S. K., and Vrijenhoek, R. C., 2004. Osedax: Bone-eating marine worms with dwarf males. Science, 305, 668–671. Smith, C. R., and Baco, A. R., 2003. Ecology of whale falls at the deep-sea floor. Oceanography and Marine Biology: An Annual Review, 41, 311–354. Smith, C. R., Kukert, H., Wheatcroft, R. A., Jumars, P. A., and Deming, J. W., 1989. Vent fauna on whale remains. Nature, 341, 27–28. Squires, R. L., Goedert, J. L., and Barnes, L. G., 1991. Whale carcasses. Nature, 349, 574. Turner, R. D., 1973. Wood-boring bivalves, opportunistic species in the deep sea. Science, 180, 1377–1379.

Cross-references Beggiatoa Cold Seeps Deep Biosphere of the Oceanic Deep Sea Hydrothermal Environments, Marine Sulfate-Reducing Bacteria

Z

ZINC

highly insoluble ZnS mineral formation, which is strongly linked to sulfur-based microbial activity.

Matthias Labrenz1, Gregory K. Druschel2 IOW-Leibniz Institute for Baltic Sea Research, Section Biology, Rostock-Warnemuende, Germany 2 University of Vermont, Burlington, VT, USA

Abundance and distribution in the environment Zinc is one of the world’s principle ores and stands fourth among all metals in world production (behind iron, aluminum, and copper), used in a variety of ways from metal products to rubber production to medicines (USGS, 2008). The world’s total reserve of this widely distributed metal is approximately 460 million tons in the year 2006 (Cohen, 2007). More than 80 zinc minerals are known; ZnS (as cubic sphalerite and hexagonal wurtzite polymorphs) and its weathering products are the most important minerals for commercial use (Henkin, 1984), with smithsonite (ZnCO3) and willemite (Zn2SiO4) being important non-sulfide forms.

1

Synonyms Spelter (nonscientific) Definition Physicochemical characteristics Zinc (Zn) is the 23rd most abundant element in the Earth’s crust, and exists as a blue-whitish metal relatively weak with a melting point of 419.5 C and a boiling point of 907 C, with a density of 7.133 g/cm3 (Henkin, 1984). Zn consists of a mixture of the five stable isotopes 64Zn (48.6%, atomic mass 63.9), 66Zn (27.9%, atomic mass 65.9), 67Zn (4.1%, atomic mass 66.9), 68Zn (18.8%, atomic mass 67.9), and 70Zn (0.6%, atomic mass 69.9) (Coplen et al., 2002). Moreover, six synthetic radioactive isotopes are known: 62Zn, 63Zn, 65Zn, 69Zn, 72Zn, and 73 Zn. The atomic number is 30. Zinc metal is highly reactive and produces various different salts, but it is only stable in water as the Zn2+ ion and associated complexes and minerals. Its sulfates and chlorides are water-soluble and its sulfides, oxides, carbonates, phosphates, silicates, as well as organic complexes are water-insoluble (Henkin, 1984). Zinc, like many other metals, also exists as dissolved metal-sulfide cluster complexes (particularly Zn3S3 and Zn4S64 ), which can exist even in oxic river waters for extended periods and account for up to 20% of the total dissolved Zn in solution (Rozan et al., 2000). Zinc as an element is not available as a metabolic component as it only exists in the Zn2+ state in water, but the strong bonds formed with dissolved sulfide ions result in

Zn ore deposition The genesis of many types of ZnS ore deposits are often attributed to purely physicochemical processes, and the distinction between abiotic and potentially biotic process is often made based on the temperature of sulfide formation and subsequent ZnS mineral formation. At low temperatures (below 150–175 C), thermochemical sulfate reduction (TSR) is very slow (Goldhaber and Orr, 1995; Ohmoto and Goldhaber, 1997; Thom and Anderson, 2008), whereas the upper temperature limit for microbial sulfate reduction overlaps with the formation of many low-temperature deposits (Ledin and Pedersen, 1996; Druschel et al., 2002). ZnS ore depositional processes without microbial influence were, for example, probably due to depositional environments outside the range of temperatures where microorganisms can survive, and TSR is sufficiently fast for significant ore formation, such as observed for the formation of high temperature hydrothermal Pb–Zn ore deposits as, e.g., described for the Rhodopian metallogenic region

Joachim Reitner & Volker Thiel (eds.), Encyclopedia of Geobiology, DOI 10.1007/978-1-4020-9212-1, # Springer Science+Business Media B.V. 2011

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(Tarkian and Breskovska, 1989; Kaiser-Rohrmeier et al., 2004). On the other hand, Mississippi Valley-Type (MVT) deposits also form from highly saline brines, between 50 C and 200 C, and span depositional environments where abiotic and biotic factors may be responsible for zinc sulfide mineralization (Bastin, 1926; Druschel et al., 2002; Thom and Anderson, 2008). Siebenthal (1915) suggested, as early as 1915, that microbial sulfate reduction contributed to the formation of low-temperature strata-bound zinc sulfide deposits. However, there is considerable controversy associated with the interpretation of the complex paragenetic sequences responsible for Zn ore deposit formation, and it is entirely possible that some deposits experience both biological and abiotic ZnS formation at different times and in different places. Trudinger et al. (1972) reviewed the feasibility of biogenic ore formation and addressed three issues: (1) the environmental limits of biogenic sulfate reduction; (2) whether the age of biological sulfate reduction was coextensive with the ages of ancient deposits; and (3) whether the rates of sulfate reduction are sufficient for ore formation. Trudinger et al. (1972) concluded that the information at the time was not adequate to address these questions. However, in the last 30 years significant progress has been made on each of these issues to better evaluate the role of microorganisms in ore deposit formation (Druschel et al., 2002; Spangenberg and Herlec, 2006). Organic geochemical and isotopic indicators of biological sulfate reduction have been recently reported from some Pb and Zn deposits (Hu et al., 1998; Bechtel et al., 1998, 1999; Spangenberg and Herlec, 2006). Moreover, modern in situ formation of pure nanocrystalline ZnS deposits at low temperature from complex groundwater solutions by SRB biofilms has been documented (Labrenz et al., 2000; Labrenz and Banfield, 2004; Moreau et al., 2004). Druschel et al. (2002) modeled the source-to-sink geochemistry of this modern ZnS-forming system and reviewed the relevance of the findings to ZnS ore deposit formation. Aggregation of those 1–5 nm sphalerite and wurtzite crystals additionally emphasizes a size-dependent stability shift in sphalerite–wurtzite stability and the role of extracellular proteins in the aggregation of ZnS nanocrystals (Zhang et al., 2003; Moreau et al., 2007). Luther et al. (1999) also showed the importance of zinc-sulfide molecular clusters (particularly Zn3S3 and Zn4S64 ) on the speciation of dissolved zinc and their importance in ZnS nanocrystal formation. Oxidation of ZnS minerals can occur abiotically in the presence of oxygen or other oxidants (Rimstidt et al., 1994), but is oxidized much faster via the activity of oxidizing microbes. Oxidative dissolution of ZnS and other metal sulfides is the principle cause of acid mine drainage (AMD), forming acidic, metal-rich waters (Nordstrom, 2000; Druschel et al., 2004). ZnS mineral oxidation releases dissolved Zn2+ ions and associated complexes into aqueous solutions. When AMD waters containing Zn2+ are neutralized, zinc can precipitate as a carbonate or oxide form, but is more commonly associated with iron oxide and oxyhydroxide minerals due to sorption of Zn2+

to those mineral surfaces, which can remove the majority of dissolved zinc from solution at circumneutral pH (Jönsson et al., 2006).

Biological effects of Zinc: Zn contamination and bioremediation As a micronutrient Zn(II) is an ubiquitous essential metal ion playing an important role in organisms throughout the three domains Bacteria, Archaea, and Eukarya. It is essential for numerous physiological processes, serves as a cofactor in members of all six major functional classes of enzymes and is especially important in the maintenance of protein structure (Blencowe and Morby, 2003). In excess it can have toxic effects, but usually Zn deficiency is more critical for most of the higher organisms (Henkin, 1984). Usually due to anthropogenic activity zinc has been shown to exist at significantly elevated levels in groundwaters and in soil and sediments (Sani et al., 2001). It has repeatedly been demonstrated by cultivationdependent as well as independent methods that this contamination can have ecotoxicological effects on, e.g., soil microorganisms, and can change the microbial community composition and activity (Hanbo et al., 2004; Smolders et al., 2004). Unfortunately, critical thresholds for toxic effects of Zn on microorganisms are not easy to develop because these can be influenced by the exposure time of Zn to the environment (Mertens et al., 2006) and the potential adaptation of microbial populations to changing Zn concentrations over time. Great interest in Zn–microbe interactions has arisen in recent years as scientists and engineers try to remove, recover, or stabilize Zn in soils, contaminated water, or waste streams (Sani et al., 2001). In general, specific metabolic pathways leading to precipitation of heavy metals as metal sulfides, phosphates, or carbonates possess significance for possible biotechnology applications (Kotrba and Ruml, 2000) and, analogous to their potential role in low-temperature Zn ore deposition, SRB are already commonly used for the bioremediation of metal-contaminated soil or water. White et al. (1998), for instance, described an integrated microbial process for the bioremediation of soil contaminated with toxic metals using microbially catalyzed reactions. In this process, bioleaching of Cd, Co, Cr, Cu, Mn, Ni, and Zn via sulfuric acid produced by sulfur-oxidizing bacteria was followed by precipitation of the leached metals as insoluble sulfides by the action of SRB. Conclusion Zinc is a common element throughout the world, mainly concentrated in zinc sulfide deposits, and has an important biological role as an essential metal ion in organisms of all three kingdoms. Though microorganisms do not directly metabolize forms of zinc, the mobility and in part the deposition of Zn2+ as ZnS minerals is strongly dependent on the activity of sulfur-reducing and sulfuroxidizing microorganisms. Indications exist for the active

ZINC

involvement of ancient sulfate-reducing bacteria in the genesis of low-temperature Zinc ore deposits and a potential biosignature capacity of nanocrystalline sphalerite for this process; however, these aspects will need further investigation.

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Kotrba, P., and Ruml, T., 2000. Bioremediation of heavy metal pollution exploiting constituents, metabolites and metabolic pathways of livings. A review. Collection of Czechoslovak Chemical Communications, 65, 1205–1247. Labrenz, M., and Banfield, J. F., 2004. Sulfate-reducing bacteria dominated biofilms that precipitate ZnS in a subsurface circum-neutral pH mine drainage system. Microbial Ecology, 47, 205–217. Labrenz, M., Druschel, G. K., Thomsen-Ebert, T., Gilbert, B., Welch, S. A., Kemner, K. M., Logan, G. A., Summons, R. E., De Stasio, G., Bond, P. L., Lai, B., Kelly, S. D., and Banfield, J. F., 2000. Sphalerite (ZnS) deposits forming in natural biofilms of sulfate-reducing bacteria. Science, 290, 1744–1747. Ledin, M., and Pedersen, K., 1996. The environmental impact of mine wastes—roles of microorganisms and their significance in treatment of mine wastes. Earth-Science Review, 41, 67–108. Luther, G. W., Theberge, S. M., and Rickard, D. T., 1999. Evidence for aqueous clusters as intermediates during zinc-sulfide formation. Geochimica et Cosmochimica Acta, 63, 3159 3169. Mertens, J., Springael, D., De Troyer, I., Cheyns, K., Wattiau, P., and Smolders, E., 2006. Long-term exposure to elevated zinc concentrations induced structural changes and zinc tolerance of the nitrifying community in soil. Environmental Microbiology, 8, 2170–2178. Moreau, J. W., Webb, R. I., and Banfield, J. F., 2004. Ultrastructure, aggregation-state, and crystal growth of biogenic nanocrystalline sphalerite and wurtzite. American Mineralogist, 89, 950–960. Moreau, J. W., Weber, P. K., Martin, M. C., Gilbert, B., Hutcheon, I. D., and Banfield, J. F., 2007. Extracellular proteins limit the dispersal of biogenic nanoparticles. Science, 316, 1600. Nordstrom, D. K., 2000. Advances in the hydrogeochemistry and microbiology of acid mine waters. International Geology Review, 42, 499–515. Ohmoto, H., and Goldhaber, M. B., 1997. Sulfur and carbon isotopes. In Barnes, H. L. (ed.), Geochemistry of Hydrothermal Ore Deposits, 3rd edn. New York: Wiley, pp. 517–611. Rimstidt, J. D., Cermak, J. A., and Gagen, P. M., 1994. Rates of reaction of galena, sphalerite, chalcopyrite, and arsenopyrite with Fe(III) in acidic solutions. In Alpers, C. N., and Blowes, D. W. (eds.), Environmental Geochemistry of Sulfide Oxidation. ACS Symposium Series 550. Washington DC: American Chemical Society, pp. 2–13. Rozan, T. F., Lassman, M. E., Ridge, D. P., and Luther, G. W., 2000. Evidence for iron, copper, and zinc complexation as multinuclear sulphide clusters in oxic rivers. Nature, 406, 879–882. Sani, R. K., Peyton, B. M., and Brown, L. T., 2001. Copper-induced inhibition of growth of Desulfovibrio desulfuricans G20: assessment of its toxicity and correlation with those of zinc and lead. Applied and Environmental Microbiology, 67, 4765–4772. Siebenthal, C. E., 1915. Origin of the lead and zinc deposits of the Joplin region. U.S. Geological Survey Bulletin, 606, 283. Smolders, E., Buekers, J., Oliver, I., and McLaughlin, M. J., 2004. Soil properties affecting microbial toxicity of zinc in laboratory-spiked and field-contaminated soils. Environmental Toxicology and Chemistry, 23, 2633–2640. Spangenberg, J. E., and Herlec, U., 2006. Hydrocarbon biomarkers in the Topla-Mezica zinc–lead deposits, northern Karavanke/ Drau Range, Slovenia: paleoenvironment at the site of ore formation. Economic Geology, 101, 997–1021. Tarkian, M., and Breskovska, V., 1989. Greenockite from the Madjarovo Pb–Zn ore district, Eastern Rhodope, Bulgaria. Mineralogy and Petrology, 40, 137–144. Thom, J., and Anderson, G. M., 2008. The role of thermochemical sulphate reduction in the origin of Mississippi Valley-type deposits. I. Experimental results. Geofluids, 8, 16–26.

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Trudinger, P. A., Lambert, I. B., and Skyring, G. W., 1972. Biogenic sulfide ores: a feasibility study. Economic Geology, 67, 1114–1127. USGS (United States Geological Survey), 2008. Zinc statistics and information. Available from: http://minerals.usgs.gov/minerals/ pubs/commodity/zinc/. White, C., Sharman, A. K., and Gadd, G. M., 1998. An integrated microbial process for the bioremediation of soil contaminated with toxic metals. Nature Biotechnology, 16, 572–575. Zhang, H., Huang, F., Gilbert, B., and Banfield, J. F., 2003. Molecular dynamics simulations, thermodynamic analysis and experimental study of phase stability of zinc sulphide nanoparticles. Journal of Physical Chemistry B, 107, 13051 13060.

Cross-references Biosignatures in Rocks Divalent Earth Alkaline Cations in Seawater Isotopes Metalloenzymes Microbial Communities, Structure, and Function Nanocrystals, Microbially Induced Sulfate-Reducing Bacteria Sulfur Cycle

Author Index

A Altenbach, Alexander V., 393 Anbar, Ariel D., 502 Andersson, Andreas J., 137 Arp, Gernot, 20 Aubrecht, Roman, 836, 847 B Beimforde, Christina, 112 Bianconi, Giovanna, 599 Boenigk, Jens, 746 Boetius, Antje, 36 Bosak, Tanja, 223 Böttcher, Michael E., 541, 859, 864 Brinkmann, Nicole, 10, 306, 326 Brocks, Jochen J., 147, 167 Büdel, Burkhard, 273, 348, 401 Bürgmann, Helmut, 575 Butler, Alison, 565 C Canganella, Francesco, 599 Canuel, Elizabeth A., 230 Cavalazzi, Barbara, 189 Chen, Zhe, 58 Cockell, Charles S., 69, 143 Cypionka, Heribert, 8, 853 D Dattagupta, Sharmishtha, 866 de Beer, Dirk, 658 Decho, Alan W., 359 Défarge, Christian, 697 Dieing, Thomas, 754 Dietzel, Martin, 261 Drake, Harold L., 1 Droser, Mary L., 886 Druschel, Gregory K., 905 Dupraz, Christophe, 617 E Ehrlich, Hermann, 796 Eisenhauer, Anton, 227, 331 Emerson, David, 535 Engel, Annette S., 521 F Fang, Jiasong, 738 Farmer, Jack D., 73 Fendrihan, Sergiu, 313

Fleissner, Gerta, 441 Fleissner, Guenther, 441 Flores, Gilberto E., 456 Föllmi, Karl B., 732 Fowle, David A., 614, 654 Friedl, Thomas, 10, 306, 326 Friedrich, Michael W., 592 Fritz, Hans-Joachim, 533 G Gadd, Geoffrey M., 416 Gehling, James G., 886 Gilichinsky, David A., 726 Gingras, Murray K., 481, 777 Gischler, Eberhard, 238 Glasauer, Susan, 681 Golubic, Stjepko, 117, 657 González-Muñoz, Maria T., 185 Gorin, Georges E., 677 Grazhdankin, Dmitriy, 342 Grice, Kliti, 147, 167 Grunau, Alexander, 355 Gudo, Michael, 441 H Hansen, Bent T., 516 Hardison, Amber K., 230 Heim, Christine, 396, 586, 754, 871 Hoefs, Jochen, 511 Hoehler, Tori M., 451 Hoffman, Paul F., 814 Hoffmann, Friederike, 840 Hoffmann, Veit-Enno, 433 Hofmann, Beda A., 691, 751, 761, 851 Homann, Vanessa V., 565 Hoppert, Michael, 81, 482, 558, 772 J Jackson, Daniel J., 53, 716 Jenkins, Robert G., 278 Jensen, Sören, 886 Jimenez-Lopez, Concepción, 185 Jones, Brian, 447, 467, 608, 808 K Kano, Akihiro, 889 Kappler, Andreas, 92, 777 Kasten, Sabine, 742 Kazmierczak, Jozef, 824, 829 Kempe, Stephan, 824, 829

Kenward, Paul A., 336 Kiel, Steffen, 901 Knauth, L. Paul, 769 Knittel, Katrin, 36 Konhauser, Kurt O., 92, 274, 481, 608, 614, 654, 777 Kothe, Erika, 596, 833 Kraemer, Stephan M., 290, 793 Krüger, Martin, 684 Küsel, Kirsten, 1 L Labrenz, Matthias, 905 Lausmaa, Jukka, 883 Lee, Natuschka M., 373 Li, Guoxiang, 58 Li, Zheng-Xiang, 206 Liebetrau, Volker, 413 Liebl, Wolfgang, 553 Litchfield, Carol D., 765 Löffler, Frank E., 373 M Mackenzie, Fred T., 137 Meert, Joseph G., 434 Meisinger, Daniela B., 373 Mohr, Kathrin I., 10, 306, 326 N Nordstrom, D. Kirk, 856 O Oremland, Ronald S., 69, 784 Oschmann, Wolfgang, 201 P Pacton, Muriel, 677 Pedersen, Karsten, 411 Perry, Randall S., 322 Petsch, Steven T., 234 Pommier, Thomas, 89 Porada, Hubertus, 547 Pósfai, Mihály, 537 Posth, Nicole R., 92 Pratt, Brian R., 662 Q Quéric, Nadia-Valérie, 317, 396, 791

910 R Radtke, Gudrun, 117, 657 Ragazzi, Eugenio, 24 Rathsack, Kristina, 317 Rawlings, Douglas Eric, 182 Reid, R.Pamela, 617 Reimer, Andreas, 20 Reitner, Joachim, 134, 136, 229, 292, 293, 317, 341, 396, 454, 543, 563, 606, 657, 666, 876 Renaut, Robin W., 447, 467, 808 Reysenbach, Anna-Louise, 456 Riding, Robert, 211, 635 Riedel, Kathrin, 355 Rivkina, Elizaveta M., 726 Roberts, Jennifer A., 336 Rodriguez-Gallego, Manuel, 185 Rodríguez-Martínez, Marta, 396, 667, 893 Rodriguez-Navarro, Carlos, 185 Rothballer, Michael, 373 Russell, Michael J., 701

AUTHOR INDEX S Schieber, Jürgen, 486, 785 Schink, Bernhard, 48 Schläppy, Marie-Lise, 840 Schmid, Michael, 373 Schmidt, Alexander R., 24 Schmidt, Kerstin, 736 Schubert, Carsten J., 578 Schulz-Vogt, Heide N., 111, 877 Sephton, Mark A., 568 Severmann, Silke, 502 Simon, Meinhard, 880 Sjövall, Peter, 883 Stan-Lotter, Helga, 313, 437 Stolz, John F., 69, 784 Straub, Kristina L., 367, 370, 412

Toporski, Jan, 754 Tribollet, Aline, 117, 657

T Thiel, Volker, 64, 271, 272, 277, 362, 686, 754, 792 Thorseth, Ingunn H., 103

Z Zhang, Li, 738 Zhu, Maoyan, 58 Zielinski, Frank, 866

V Verrecchia, Eric P., 721 Visscher, Pieter T., 617 Vortisch, Walter, 266 W Warren, Lesley A., 5 Weber, Bettina, 348, 401 Westall, Frances, 189 Wiese, Frank, 293 Wörheide, Gert, 53 Wrede, Christoph, 482

Subject Index

A Abietane, in resins, 27 Abietic acid, in resins, 26 Abrasion, of biofilms, 325 Absolute age, 415 Acantharia, 334 Acanthodendrilla, 801 Acarospora, 403 A. rugulosa, 424, 425 A. smargdula, 424, 425 Accumulation rates, 778 Acetabularia, 243 Acetate, 282, 703, 779 fermentation, 282 fermentation, biomarkers, 174 synthesis of, 1 Acetic acid, 292 Acetitomaculum, 2 Acetoanaerobium, 2 Acetobacterium, 2, 86 A. woodii, 83 Acetoclastic methanogenesis, 66, 872 Acetogenesis, 1, 702 Acetogenic bacteria, 1, 86, 872 Acetogenium, 86 Acetohalobium, 2 Acetone, 292 Acetonema, 2 Acetyl CoA synthethase/carbon monoxide dehydrogenase, 559 coenzyme-A (Acetyl CoA), 559, 598, 703 pathway, 1, 705 synthase, 684 phosphate, 706 Acharax, 285 Achnanthidium saprophila, 328 Achromatium, 878 Acidianus, 184 A. brierleyi, 368, 857 Acidic ocean, 703 Acidic organic macromolecules (AOM), 673 Acidic springs, 469 Acidification, 228, 353 of the ocean, 137, 545 by fungi, 423 Acid-insoluble minerals, 183

Acidiphilium A. cryptum, 371, 372 A. rubrum, 370 Acidithiobacillus, 6, 87, 184, 362, 878 A. ferrooxidans, 6, 368, 372, 433, 856 A. thiooxidans, 433 Acid mine drainage, 5, 856, 878 Acidobacterium, 84 Acidophiles, 362, 460 iron oxidizing, 368 iron-reducers, 372 Acidophilic methanotrophs, 576 Acidovorax, 369 Acid rain, 112 Acid rock drainage, 5, 750 Acid-soluble minerals, 183 Aciduliprofundum, 462 Acid-volatile iron monosulfides (AVS), 487 Acritarchs, 202, 341, 750 Acropora, 245 Actinide, 751 Actinobacteria, 86, 90 Actinomycetes, 151, 667 Adenosine diphosphate (ADP), 706 Adenosine phophosulfate, 854 Adenosine triphosphate (ATP), 596, 706 Adsorption, of organic matter on minerals, 235 Aerobactin, 566 Aerobes, 460 Aerobic biofacies, 203 respiration, 8, 575 metabolism, 8 Agarose, 12 Agate, 272 Age, 413 absolute, 413 determination, 413 relative, 413 Aglaophyton, 301 Agronomic revolution, Cambrian, 298 Akilia, 191 Akinetes, in cyanobacteria, 307 Alabandite, 541 Alcaloids, 597 Alcolapia grahami, 477 Aldanella, 62 Algae, 10, 348 biomarkers, 171

endolithic, 349 endoliths, 122 eukaryotic, 10, 331, 441 role in bioerosion, 118 Algaenan, 157 Aliphatic molecules, 571 Alkali metals, 457 in marine hydrothermal environments, 457 Alkaline hydrothermal vent, 704 Alkalinity, 7, 20, 67, 139, 829 engine, 625 pump, 827, 828 Alkaliphiles, 362, 767, 831, 878 Alkaliphilic lakes, 362, 765, 824 Alkalization, 353, 407 n-Alkanes, 157, 158, 171, 235 Alkenones, 173, 235 Alkyl phosphonic acids, 572 Alkyl sulfonic acids, 572 Allochems, 894 Allochromatium, 602 Allomicrites, 667, 893 Alpha-carbonic anhydrase, 55 Alpha-proteobacteria, 83, 90, 539 Alteration, basalt, glass, 103, 107 Alternaria, 402, 423 A. alternata, 424 Alterobactins, 566 Aluminosilicates, 144 formation during weathering, 144 Alunite, 448 Alveolata, 747 Alveolates, 10 Alvinella, 463 Amber, 24 chemical composition, 29 Amides, in meteorites, 572 Amines, in meteorites 572 Amino acids, 235 in meteorites, 570 first synthesis, 707 chirality, 707 Ammonia (NH3), 685, 686, 704 assimilation, 687 at cold seeps, 282 Ammonifex degensii, 109 Ammonification, 49, 673, 687 Ammonium (NH4+), 577, 686, 779 Ammonium-oxidizing bacteria, 779

912 Amoebae, 393, 747 Amorphous calcium carbonate (ACC), 54, 261 Amphibactins, 565 Amphora, 329 Amylase, 357 Amyloid, 708 proteins, 360 Anabaena, 476, 655, 812 Anabarites, 62 Anaerobes, 460 Anaerobic biofacies, 203 Anaerobic degradation, 48, 236, 578, 779 Anaerobic metabolism, 48 energetics, 50 phototrophic, 51 Anaerobic oxidation of methane, 36, 224, 283, 579 role in carbonate precipitation, 223 Anaerobic methanotrophic archaea (ANME), 36 Anaerobic respiratory pathways, 48, 779 Anammox, 688, 779 Anammoxosome, 85 Anatase, 191 Ancient Archaeal Group, 65 Andisols, 144 Anema, 403 Angiosperms, biomarkers, 158, 173 Angulocellularia, 214, 215 Angusticellularia, 214, 215, 218, 643 Anhydrite, in hydrothermal environments, 457 Animals, 463, 716, 782 biocalcification, 53 skeletons, 58 Ankerite, 92, 262, 721, 789 ANME archaea, 36, 66, 463, 607, 885 Anomalocaris, 300 Anoxic basins, organic matter degradation, 236 events, cause for mass extinctions, 302 zone, in sediments, 780 Anoxygenic photosynthesis, 692, 738 in sulfur bacteria 878 Antarctica endoliths from, 348 saline lake, 765 ice, 568 Anthropogenic contaminations, 598 Antibodies, 482 Antigens, 482 Anti-Stokes scattering, 755 Apatite, 60, 145, 191, 512 early animal skeletons, 60 volcanic rock weathering, 145 Apex cherts, 95 Aphanothece halophytica, 438 Aphrocallistes vastus, 802 Aphyllophorales, 425 Apicomplexa, 10, 16 Aplysina aerophoba, 843 Apusomonads, 747 Aquabacterium, 315 Aquachelins, 565 Aquaculture, 768 Aquatic sediments, 742 Aquifers, 598 Aquifex, 82, 83, 364, 448, 476 A. pyrophilus, 448 Aquificales, 83, 461, 612 Aragonite, 60, 239, 261, 459, 836 at cold seeps, 284 compensation depth, 258 at hot springs, 448 metal ions in, 261 Araphid diatoms, nonmotile, 326 Araucaria columnaris, 32

SUBJECT INDEX Arbuscular mycorrhizal fungi, 867 Archaea, 282, 437, 448, 476, 578, 711, 835 biomarkers, 158 halophilic, 314 iron-oxidizing, 367 in marine hydrothermal environments, 461 role in dolomite formation, 337 Archaean, 202, 294 Ocean, 94 Archaebacteria. See Archaea Archaeocyatha, 59 Archaeoglobales, 368, 462 Archaeoglobus, 87, 109, 319, 854 A. fulgidus, 2 Archaeol, 37, 175 Archaeospira, 62 Archiasterella, 61 Arcobacter, 879 A. sulfidicus, 879 Arid (dry) environments, 365 Aromatic hydrocarbons, in meteorites, 571 degradation of, 598 Arsenic (As), 69, 448, 691 Artemia, 362 A. salina, 365 Arthrobacter, 359 Arthropods, preservation in amber, 33 Arthrospira, 309 Artifacts, mistaken as biosignatures, 195 Aspartic acid, role in biomineralization, 54 Aspergillus, 359, 423 A. niger, 424, 425 Aspicilia A. alpina, 424, 425 A. calcarea, 425 A. flavida, 350 A. radiosa, 425 Assimilative sulfate reduction, 859 Asteroid, 293, 568 belt, 568 Astrobiology, 73 Astrosclera willeyana, 55 Atar group, 664 Athalassohaline, 437, 765 Atmosphere, 577 oxygenation, 296 Atmospheric CO2, 732 Atomic force microscopy (AFM), 360, 772 ATP, 49, 50, 575 yield, of aerobic respiration, 8 ATPase, 575 Attachment, 359 Authigenic clay minerals, 552, 785 mineral formation, 782 quartz (chalcedony), 789 theory, 702 Automicrites, 258, 667, 893 Autotrophy, 856 Avalon-type biota, 345 Azoarcus, 598 Azolla, 309 Azotobacter vinelandii, 795 Azurite, 262 B Bacillariales, 881 Bacillariophyta, 326, 747, 881 Bacillus, 85, 86, 319, 359, 604, 687, 872 B. alcalophilus, 362 B. cereus, 187 B. infernus, 872 B. lentus, 187

B. pasteurii, 187 B. sphaericus, 187 B. subtilis, 187, 276, 433, 612, 615, 655 Bacinella, 644, 671 Bacteria, 31, 81, 348, 448, 578, 809, 810, 835 planktonic, 89 biomarkers, 159 in marine hydrothermal environments, 461 Bacteriastrum, 329 Bacteriochlorophyll, 83, 172 Bacterioferritin, 560 Bacteriohopanepolyols, 159 Bacteriomorphs, 195 Bacteriophages, 767, 873 Bacterioplankton, 89 Bacteriorhodopsin, 65 Bacteriosponges, 840 Bacteroides, 84 B. ruminicola, 84 Bacteroidetes, 90 Baeocytes, 307 Baeomyces, 402 Bahamas, 617, 243 stromatolites, 246 Baicalia, 640 B. lacera, 217 Bajanophyton, 215 Banded iron formations (BIFs), 92, 196, 294, 369, 505, 692, 817 Bangiomorpha pubescens, 12 Barberton, 195 Barite, 191, 817 precipitation by foraminifera, 394 Barium, at cold seeps, 282 Barophilic bacteria, 738 Basalt, 103, 457 weathering, 144 caves, 526 hydrothermal fluids, 457 Basidiomyces, 359 Basidiomycete, 359, 598 Bathymodiolian mussels, 287, 902 Bathymodiolus, 255, 284, 463 Batillaria, 243 Batinivia, 215 Batophora, 243, 620 Beaches, carbonate deposits, 241 Beauveria caledonica, 424, 425 Beggiatoa, 42, 87, 111, 194, 272, 283, 319, 604, 838, 859, 877, 902 B. alba, 83 Belt supergroup, 663 Benthos, 782 Benzene, 598 Benzothiophenes, 572 Benzoyl CoA, 598 Beta-carotene, 10, 151 Beta-proteobacteria, 83, 90 Bicarbonate, 20, 333, 829 waters, 470 Bicosoecida, 747 Big Five, mass extinction events, 301, 543 Bija, 215 Bilateromorpha, 343 Bioabrasion, 117 Bioalteration, basalt, 103 Bioastronomy, 73 Biocalcification advent, 58, 296 animals, evolution of, 53 structural genes, 54 Biocides, 115 Biocoenosis, 33 Bioconservation, 185

SUBJECT INDEX Bioconsolidation, 185 Biocorrosion, 117, 657 Biodeformation, 481 Biodeterioration, of building stones, 112 by fungi, 418 vs. bioprotection, 186 Biodiversity, 768 Bioelements, as biosignatures, 190 Bioeroding organisms, biodiversity, 120 Bioerosion, 117 in coral reefs, 129 over geological time, 124 phanerozoic, 125 predators in, 125 proterozoic, 124 rates of, 128 taphonomy, 124 Bioerosional notch, 127 Biofilms, 112, 120, 134, 329, 350, 359, 699, 700 biodeterioration, 112 fossilization, 136 role in carbonate precipitation, 224 Biogenicity, biosignatures, 197 Biogeochemical cycling, 420 Bioirrigation, 138, 482, 742 Biokarst, 127 Bioleaching, 182 Biological elements, 76, 686 Biologically controlled mineralization, 53, 54, 192, 586, 683 Biologically induced mineralization, 53, 587, 681 Biological soil crust (BSC), 403 Biomarkers, 147, 189, 441 biological sources of, 153 compound-specific isotopes, 167 of cyanobacteria, 170 of methane oxidizing bacteria, 175 of methanogenic archaea and methane oxidizing archaea, 175 of phytoplankton, 170 of zooplankton, 170 of diatoms, 156 different meanings of, 148 of eukaryotes, 152 of green sulfur bacteria, 160 molecular fossils, 331, 441 precambrian, 96 of sponges, 158 Biomimetic materials, 56 Biomineralization, 300, 698, 700, 830, 837 animals, 53, 58 by Archaea, 66, 336 basalt, 106 clay, 275 of fungi, 424 inhibition, 263 macromolecules, 54 microbial, 586, 608 vs. organomineralization, 698 sponges, 796 Biominerals, 618, 587 vs. organominerals, 697 as biosignatures, 191 formation, 773 Biomining, 182 Biooxidation, 182, 691, 856 Biopatina, 113 Biopolymers, in soil, 235 Biopreservation, 185 Bioprotection, 185 by endoliths, 352

Biopsy, 441 Bioreactors, 7 Bioremediation, 413, 598, 906 by fungi, 420 Biorestoration, 185 Biosignatures, 189, 352, 700 astrobiological implications, 196 metal isotope fractionation, 503 in microbial dolomite, 339 isotopes, 511 molecular (biomarkers), 147, 167 of microbially induced carbonates, 224 Biosilicification, 608, 796 Biosphere, 732 definition, 313, 317 Biostabilization, 548 Bioturbation, 201, 481, 742, 782, 886 control on diagenesis, 138 Bioweathering, 143, 422 Biphytane, 158, 175 Bird, 767 Birnessite, 424 Bitumen, 150 Bituminous mudstones, 202 Bivalves, at cold seeps, 278, 287 Bjerkandera, 420 Black carbon, 235 Black crusts, 185 Black fungi, 402 Black holes, 599 Black Sea, 42 Black shales, 201, 295, 785 Black smokers, 318, 454, 458 Bleaching, 16, 123 Blue-green algae. See Cyanobacteria Blue holes, 599 Bodonids, 747 Bones, Sr-isotopes, 517 Boring, 117, 146 foraminifera, 123 sponges, 119 traces, 121 Borrelia, 84 Bositra, 398 Botomaella, 643 Botominella, 215 Botryococcus, 157, 171 Botryococcus braunii, 153, 157, 171 Brachiopods, at cold seeps, 287 Bradoriida, 59 Braziliensis, 749 Brevibacterium, biomarkers, 151 Brevibacterium linens, 151 Brine flies, 767 shrimp, 767 springs, 767 Brocadia anammoxidans, 85 Brown algae, 747 Brown rot fungi, 597 Brucite (Mg (OH)2), 459 Bryantella, 2 Buellia, 348, 403 Building stones, biodeterioration, 112 Burgess Shale, 296, 785 Burial efficiency, 236 Burkholderia, 145, 359 Burndown, 496, 498 Burrowing, 117, 481, 782 Burrowing organisms, 481, 782 Burrow linings, 481, 782 Butyribacterium, 2 Byroniida, 59

913 C

14

C, 415 Ca2+-ATPase, 55 Cadmium (Cd), 424, 448 Calcarea, 59, 797 Calcareous algae, 211, 224 Calcareous oozes, 278 Calcification, 13, 211, 441, 588, 698 animals, 53 cyanobacteria, 211, 309 impact on alkalinity, 23 microbial, 223 microbialites, 625, 636 Calcified microfossils, ancient, 196 Calcimicrobial reefs, 218 Calcite, 12, 60, 261, 309, 424, 785 accumulation in algae, 12 budget, 394 compensation depths, 258 at hot springs, 448 metal ions in, 261 microbialites, 636 microbial precipitation, 223 Calcium (Ca), 54 detoxification, 54 in seawater, 332 isotopes, 227, 264, 335 isotope thermometry, 335 oxalate, 144, 722 phosphate, as biomineral, 54 rhodochrosites, 541 ions in seawater, 331 Calcium carbonate, 54, 261, 264, 698, 700 CaCO3 polymorph, 262 CaCO3 precipitation, by cyanobacteria, 211 CaCO3 supersaturation, 23 nucleation on organic surfaces, 224 Calcretes, 721, 724 Caldera lakes, 827 Caldivirga, 854 Callianassa, 243, 248 Callisto, Jupiter’s moon, 77 Calmodulin, 54 Caloplaca, 403, 407 C. aurantia, 424, 425 C. citrine, 402 C. flavescens, 425 Caloramator, 2 Calothrix, 135, 476, 609, 812 Calvi, 141 Calvin cycle, 172, 737 Calyptogena, 42, 255, 281, 285, 463, 879 Cambrian boundary, 733 Cambrian Evolutionary Fauna, 63 Cambrian explosion, 53, 62, 296, 663, 718, 771 Cambrian radiation, 58, 62, 218 Cambroclaves, 59 Cambroclavus, 61 Campbellrand Malmani platform, 646 Candelariella vitellina, 402 Canfield Ocean, 299, 543 Canopoconus, 62 Cap carbonates, 229, 287, 646, 815 Capnodiales, 402 Carbohydrates, 235, 701 oxidation of, 48 Carbon assimilation, 576 cycle, 732 isotopes, 511 d-value, 579 BIF, 98 biomarkers, 167

914 Carbon (Continued ) graphite, 512 methane, 282, 579 organic, cycling, 230 organic, degradation, 234 Carbonaceous chondrites, 273, 293, 568 Carbonate, 92, 261, 282, 352, 360, 426, 635, 829 alkalinity, 20, 626 budget, 129 compensation depth, 262 diagenesis, 138 dissolution, 129, 138, 262 bioerosion, 119 by endoliths, 129 dumbbells, 339 environments, 238, 762 fluoroapatite, 191 growth kinetics, 263 at hot springs, 470 hydroxylapatite, 191 karst, 521 marine, bioerosion, 121 mud, ancient, 216 platform, 733 pedogenic, 721 precipitation, 20, 22 diagenetic, 140 inhibition, 263 microbial, 211, 223 mediated by EPS, 360 role of alkalinity, 20 production and foraminifera, 394 reduction, 282 system, 20 Carbonatite, 824 Carbon dioxide (CO2), 278, 282, 406, 575, 702, 706, 777 atmospheric, geological record, 215 concentrating mechanisms, by phototrophs, 212 reduction, 174 Carbonic acid weathering, 829 Carbonic anhydrase, 55 biocorrosion, 119 Carboxylic acids, 570 CARD-FISH, 382 Carotenoids, 10, 151 Caspian Sea, 765 Catabolism, 596 Catagenesis, 150, 230 Catalyzed reporter deposition (CARD), 484 Cathodoluminescence Microscopy, 266 Cathepsin L, 800 Caulerpales, 11 Caulobacter, 87 Cave paintings, 528 Caves, 225, 521, 599 Cayeuxia, 215 Celestine, 191 Cell counts, bacterioplankton, 89 Cellulase, 357 Cellulose, 235, 597 degradation of, 356 Cell wall composition, role in metal fixation, 195 Cementation, 777, 782 Central Atlantic Magmatic Province (CAMP), 545 Centric diatoms, 326 Centropyxis hirsute, 31 Ceratodictyon spongiosum, 13 Cercomonas, 749 Cercozoa, 747 Cerium (Ce), 831 Chabakovia, 215 Chaetoceros, 329 Chaetognatha, 59

SUBJECT INDEX Chaetomium, 402 Chaetothyriales, 402 Chalcedony, 812 Chalcocite, 183 Chalcopyrite (CuFeS2), 191, 458, 856 Chalk, 258, 277 Challenger Deep, 317 Chamberlain formation, 664 Chamosite, 191, 275 Chara, 12, 157 Charales, 12 Charnia masoni, 346 Charniodiscus concentricus, 344 Charophyte, 12 Chasmoendolithic, 349 Chasmoendoliths, 118, 348 Chattanooga Shale, 789 Chemical force microscopy (CFM), 775 Chemical lamination, 414 Chemiosmosis, 704, 708 Chemodenitrification, 687 Chemoheterotrophy, 596, 777, 782, 838 Chemolithoautotrophy, 87, 113, 271, 463, 476, 526, 558 bacteria, 779 at cold seeps, 284 Chemoorganotrophy, 114, 476 biodeterioration, 114 Chemosymbiosis, 285 Chemosynthesis, 230, 869 at cold seeps, 278, 284 Chengjiang, 296 Chernobyl, 423 Chert, 272 Chicxulub crater, 546 Chirality, 75, 189, 570 Chitin, 235 degradation of, 356 Chitinase, 357 Chitinozoa, 750 Chlamydobacteria, 564 Chlamydomonas nivalis, 365 Chlorarachniophytes, 10, 16 Chlorella, 10, 12 Chloride-bicarbonate waters, 470 Chloride waters, 468–469 Chlorobactene, 160, 171 Chlorobiaceae, 151, 160, 171 Chlorobionts, 401 Chlorobium, 83 C. ferrooxidans, 369 Chloroflexus, 83, 90, 476, 477, 812 C. aurantiacus, 198 Chlorophyceae, 11, 12 Chlorophyll, 10, 148, 170, 307, 327, 560 Chlorophyta, 10, 12 Chlorosis, 793 Chloroxybacteria, 306 Choanoflagellates, 747, 840 Choanomonada, 747 Cholestane, 152 Chondrites, 258, 482, 568 Chondrosia reniformis, 843 Chondrules, 568 Chondrus, 12 CH4. See methane Chromalveolates, 10, 13, 15 Chromatiaceae, 151, 161, 172, 602 Chromatiales, 161 Chromatium, 83 Chromist, 750 Chromista, 746, 749 Chromohalobacter, 438 Chroococcales, 310

Chroococcidiopsis, 348, 350 Chrysolaminarin, 327 Chrysomonads, 747, 749 Chrysophytes, 14, 747 Chuaria, 341 Ciliates, 747 biomarkers of, 157, 173 C. See carbon Citrate cycle, 596 Citrobacter, 604 Cladocora, 253 Cladogirvanella, 215 Cladonia, 402 Cladonia convolute, 406 Cladophora, 10 Cladosporium, 402, 423, 424 Cladosporium cladosporoides, 424 Clathrates, 583 Clausthalite, 784 Clay, 145, 785, 833 bacterial authigenesis, 274 mineral formation, 427 minerals, 448, 785 Climate change, Nd isotopes, 518 Climate, impact of methane, 287 Cliona vestifica, 120 Clionid, 124 Clone libraries, 593 Clostridium, 2, 86, 359, 597 C. aceticum, 1, 84 C. acetobutylicum, 84 C. butyricum, 84 C. glycolicum, 3 C. klyuveri, 84 C. ljungdahli, 2 C. oroticum, 84 C. pasteurianum, 371 C. sticklandii, 84 C. tetanomorphum, 84 C. thermoaceticum, 1 Clot, 641 Clotted fabrics, 190, 196 Cloudina, 58, 60, 297 C–4 methylsteranes, 156 C:N ratio, 779 Coal, 231, 597 Cobalt (Co), 183, 424, 562 Coccales, 462 Coccolithophorids, 15, 277 Coccoliths, 15, 277 CO dehydrogenase, 684 Codon, 710 Coeloscleritophora, 59 Coenogonium, 402 CO2, fixation by acetogens, 1 Cold seeps, 42, 255, 278, 319, 576, 902 Cold-water corals, 253 Cold-water reefs and mounds, 253 Coleoloida, 59 Collagen, 802 Collema, 401, 405 Colorless sulfur bacteria, 878 Colwellia, 739 Cometabolic degradation, 597 Comets, 290, 293, 573, 701 impacts, 69 Commensalism, 290, 463, 866 Communic acid, in resins, 27 Community, microbial, 592 Compaction, 787 Compatible solutes, 364 Compound-specific isotope analysis (CSIA), 167 Concentration-depth profiles, 742 Conchocelis, 118

SUBJECT INDEX Concrete, biodeterioration by fungi, 423 Confocal Raman microscopy, 754 Confocal scanning laser microscopy (CSLM), 360 Conifer resins, 158 Coniosporium, 402 Conophyton, 640 Conotheca, 62 Consolidation treatments, 115 Contamination, biosignatures, 194 Continental deep biosphere, 871 Continental rift, 206 Continental shelves, 206 Continental weathering, 830 Conulariida, 59 Copal, 28 Copper (Cu), 183, 290, 448, 457, 575, 856 in marine hydrothermal environments, 457 removal from acid mine drainage, 7 Copper-containing proteins, 561 Coral bleaching, 123 Coralline algae, 12 Corallistes undulatus, 798 Corallistidae, 798 Coral reefs, 245, 762 bioerosion, 117, 129 CO2. See Carbon dioxide Coriolopsis, 420 Cosmic dust, 701 Cosmic molecular clouds, 292 Covellite, 183 C3-plants, 512 biomarkers, 173 carbon isotope fractionation, 512 C4-plants, 173, 512 biomarkers, 173 carbon isotope fractionation, 512 C:P ratio, 733 Crassostrea, 267 Craters, 70 Creek, 257 Crenarchaeota, 65, 158, 462, 607, 784 biomarkers, 174 cell walls, 195 Crenothrix, 529 Cretaceous, 733, 872 Cretaceous–Eocene, 733 Cretaceous/Tertiary, extinction event, 70, 302 Cristobalite, 272 Critical Intervals in earth History, 293 Crocetane, 37, 159, 175 Crookesite, 784 Crude oil, biomarkers, 148 Crustaceans, 767 Cryobiosphere, 306, 726, 364 Cryochron, 814 Cryogelling, 812 Cryogenian, 814 Cryogenian Snowball Earth, 543, 814 Cryolithosphere, 726 Cryomyces, 402, 404 Cryopegs, 728 Cryosphere, 726 Cryptalgal, 636 Cryptic environment, 838 Cryptococcus, 123 Cryptoendoliths, 118, 348 Cryptomicrobial, 636 Cryptomonads, 10, 16, 747 Cryptophyceae, 747 Cryptozoon, 638 Crystal lattice, calcite, 262 Crystalline shapes, biosignatures, 193 Cultivation, 592

Cultural heritage, 112 conservation, 185 Cunninghamella, 420 Cupithecids, 59 Cupittheca, 62 Cupriavidus (Ralstonia) metallidurans, 434 Cutan, 235 Cyanidium, 350, 477, 812 C. caldarium, 362, 476 Cyanobacteria, 83, 85, 90, 224, 231, 306, 401, 736, 810, 827, 838 ancient, 196 biomarkers, 160, 175 biomarkers (compound-specific isotopes) of, 170 calcified, 211 coccoid, 548 endolithic, 121, 349 filaments, 698 first occurrence, 95 fossilized, 215 mats, 476 in microbialites, 620, 636 role in bioerosion, 118 sheaths, 699 Cyanobionts, 401 Cyanodecapentayne, 292 Cyanophages, 307 Cyanophycin, 307 Cyanophyta. See Cyanobacteria Cyanoprokaryota. See Cyanobacteria Cyanosaccus piriformis, 119 Cyanotoxins, 309 Cyclomedusa davidi, 344 Cymbella, 328 Cymodocea, 243 Cysteamine, 876 Cysteine glycoconjugate, 535 Cystocoleus, 402 Cytochrome, 558 C, 575 oxidases, 560 P450, 559 Cytophaga, 84 D Dabashanella, 62 Damselfish, 126 DAPI, 376 Dasycladales, 11 Dating, 413 Pb isotopes, 518 Dead Sea, 765 Dead zones, 733 Deccan trap basalts, 546 role in weathering, 144 Decomposition of organic matter, 234, microbial, 596 during diagenesis, 777 Deep biosphere, 760 basalt, oceanic crust, 106 deep sea, 317 definition, 317 disruption by impacts, 72 filamentous fabrics, 851 hydrogen-driven, 872 of salt deposits, 313 terrestrial, 871 Deep Sea Drilling Project (DSDP), 318, 733 Deep-sea Hydrothermal Vent Euryarchaeotal Group, 319 Deep-sea hydrothermal vents, 456, 554, 693 Deep-water carbonates, 258 Deep-water reefs and mounds, 253

915 Deferribacter thermophilus, 371 Deformation, molar tooth structures, 662, 664, 666 Degradation, 112, 596 aerobic, 8 anaerobic, 50 by fungi, 420 microbial, 596 organic matter, 234, 777 Degree of pyritization (DOP), 493, 781 Dehalogenation, 598 Dehalorespiration, 596, 598 Deinocci, 69 Deinococcus, 84 D. radiodurans, 365, 752 Deltaproteobacteria, 83, 540, 854 Delta (d) value, 168, 511 Demospongiae, 59, 797, 840 Dendrina, 125 Dendrochronology, 414 Dendrolites, 617 Denitrification, 236, 322, 394, 687, 779 coupled with methane oxidation, 43 nitrogen isotope fractionation, 513 Depolymerization, 743 Deprotonation, of organic functional groups, 614 Dermatiscum, 403 Deserts, 350 Desert varnish, 322, 424 Desmas, 800 Desmochloris, 10, 12 Desulfobacter, 854, 859 Desulfobium, 854 Desulfobulbus, 37, 40, 854 Desulfocapsa, 859 Desulfococcus, 39, 319 Desulfonema, 604 Desulfosarcina, 37, 319, 337, 602 Desulfostipes saporovans, 337 Desulfotomaculum, 109, 854 Desulfovibrio, 854, 859 D. desulfuricans, 139, 186 D. magneticus, 540 D. vulgaris, 83, 186 Desulfuration, 859 Desulfurococcales, 462 Desulfurococcus, 319 Desulfuromonas D. acetexigens, 371 D. acetoxidans, 83 D. palmitatis, 372 Desulfuromusa kysingii, 371 Detachment, of microbial cells, 325 Deterioration, 422 Detoxification, 681, 832 Detrital iron, 781 Deuterium, in lipid biomarkers, 167 Deuterostomia, biomineralization, 56 Devonian Plant Hypothesis, 544 Dewatering, 665 Dextrorotatory, 570 D/H ratios, in lipid biomarkers, 171 Diagenesis, 137, 149, 230, 812 in BIFs, 99 organic matter decomposition, 230, 777 biological control on, 777 equation, 138 phosphate formation, 788 pore water recations, 742 redox zonation, 743 reaction-transport models, 743 silica (quartz) precipitation, 788 Diatomaceous earth, 326 Diatomite, 272

916 Diatoms, 326, 476, 747, 788, 797, 812, 838 biomarkers, 156 fossil record, 329 reproduction, 327 Diazotrophy (nitrogen fixation), 307, 686 Dicentrarchus, 767 Dickinsonia, 886 Dickinsoniomorpha, 343 Dictyochophyceae, 14 Dictyostelids, 748 Diethyl ether, in cosmic molecular clouds, 292 Diffusive seeps, 278 Digitothyrea, 403 Dikelets, 665 Dimelaena, 403 Dinitrogen, 686 Dinoflagellates, 10, 15, 331, 747 biomarkers, 156, 172 Dinosterane, 156 Di-oxygenases, 598 Diploneis, 329 Diploria, 245 Diploschistes muscorum, 406 Discharge apron, of hot springs, 809 Discoidal structures, Ediacaran biota, 345 Dismutation, of organic compounds, 50 Disproportionation, of sulfur species, 878 Dissimilatory sulfate reduction, 781, 854, 859 Dissimilatory manganese reduction, 780 Dissolution, 129 of carbonates, 262 microbial, basalt, 106 rates, 263 Dissolved inorganic carbon (DIC), 224, 262 Dissolved organic carbon (DOC), 231 Diterpenes, in resins, 26 Diurnal cycles, 51 Diversity, 336, 592 DNA, 32, 146, 709 in extracellular polymeric substances, 360 Docosahexaenoic acid, 740 Dolomite, 92, 239, 261, 331, 721, 830 at cold seeps, 284 microbial formation of, 336 problem, 337 in shales, 785 in soils, 722 Dolomitization, 336, 782 Domains of life, 64 Doratomyces, 424, 425 Dormant cells, cyanobacteria, 307 Dothideomycetes, 402 Doushantuo Formation, 788, 877 Dripstones, 836 Dumbbells, carbonate, 339 Dunaliella, 174, 767 biomarkers, 174 D. acidophila, 362 D. salina, 438 Dwarf bacteria, 678 Dykelets, 666 Dynamic SIMS, 883 Dysaerobic biofacies, 203 Dysidea avara, 843 E Early animals, skeletons, 58 Early Cambrian, skeletal metazoans, 62 Early history of the Earth and life, 293, 703 Early life, C-isotopes, 512 origin of, 701 Early ocean, 769, 829 Ebullient springs, 447

SUBJECT INDEX Ecdysozoa, biomineralization, 56 Echinodermata, 59 Echinometra mathaei, 127 Ecological amplitude, 402 Ecological niche, 592 Ecosystem, 592 Ectosymbiosis, 123, 866, 879 Ectothiorhodospira halochloris, 438 Ediacaran, 58, 297, 342, 717, 815, 830 Eel River Basin, 42 Eicosapentaenoic acid, 740 Electron acceptors, 48, 234, 596 in photosynthesis, 736 in pore waters, 742 in diagenesis, 777 used by acetogens, 3 Electron carriers, 51 Electron donors, 48 iron reduction, 371 Electron-nuclear double resonance (ENDOR) spectroscopy, 558 Electron paramagnetic resonance (EPR), 558 Electron shuttles, 780 Electron spin resonance (ESR), 558 Electron transfer, extracellular, 51 Electron transport chain, 51 Electron transport phosphorylation, 596 Electrostatic force microscopy (EFM), 775 Embden–Meyerhof pathway, 596 Embedding, of geobiological specimens, 443 Embryos, 342, 877 Emiliania huxleyi, 278 Enamel, Sr-isotopes, 517 Enceladus, Saturn’s moon, 77, 726 Encrustation, 449 by iron sulfides, 492 of microbes in sinter, 449 End-Cretaceous mass extinction, 302, 545 Endoflagellum, 84 Endoliths, 71, 103, 348 algae, 118 in basalt, 108 bioerosion, 118 borings, 145 in corals, 123 cyanobacteria, 273 fungi, 118 in glass, 103 growth, 402 microbial, evolution, 124 volcanic rock weathering, 145 Endomycopsis fibuligera, 433 Endomycorrhiza, 867 End-Ordovician mass extinction, 301, 543 Endoskeletons, 58 Endospores, 86, 307 Endosymbiosis, 10, 355, 394, 463, 576, 763, 866, 879 hypothesis, 867 End-Permian mass extinction, 301, 545 biomarkers, 172 End-Triassic mass extinction, 302, 545 Energetics, of anaerobic metabolism, 50 Energy conservation, 596 anaerobic, 48 Energy sources, classification of, 459 Energy yield, metabolism, 778 Engineering applications, 700 Entamoeba histolytica, 746 Entermorpha prolifera, 172 Enterobacter, 359 E. aerogenes, 84 Enteromorpha, 157 Entner–Doudoroff pathway, 596

Entophysalis, 643 E. major, 350 Enzyme sharing, 395 Eocene, 733 Eohardrotreta, 62 Eohyella, 353 Eosphaera tyleri, 341 Ephebe, 402 Ephemeral lake, 765 Ephydatia fluviatilis, 799 Ephydra, 362, 477 Epibionts, 879 Epicuticular waxes, biomarkers, 173 Epiliths, 118 Epiphyton, 215, 218, 399, 643, 671 Episymbiosis, 463. See Ectosymbiosis Epitope, 483 EPS. See Extracellular polymeric substances (EPS) Epsilon-proteobacteria, 83, 879 Equilibrium effects, isotopes, 167, 512 Ergostane, 152 Ernietta, 298 Erosion, of biofilms, 325 Escherichia, 371, 604 E. coli, 69, 81, 84, 85, 433, 533, 556, 655 Etching, 144 Ethane, 282 Ether lipids, archaeal, 158 Ethyl cyanide, 292 Eubacteria, 2 Euglenids, 10, 17, 747 Eugomontia, 121 Eukaryota, 204, 453, 831, 835 biomarkers of, 152 early precambrian, 341 Eulitoral, 880 Euphotic zone, 122 Euplectella aspergillum, 800 Europa, Jupiter’s moon, 77, 726 Eurotiomycetes, 402 Euryarchaeota, 37, 65, 282, 462, 687 cell walls, 195 Eutrophication, 203, 733 Euxinic waters, 172 Evaporites, microbes in salt deposits, 313 halophiles, 437 hypersaline environments, 362 saline lakes, 765 soda lakes, 824 stromatolites, 648 Everything is everywhere, 750 Evolution, driving force of predation, 54 Evolutionary tree, 711 Excavates, 10 Excessive mucilage events, 361 Excrements, of birds, 112 Exfoliation, 353, 407 Exobiology, 73 Exoenzymes, 355, 778 Exopolymers, See Extracellular polymeric substances (EPS) Exopolysaccharides, See Extracellular polymeric substances (EPS) Exoskeleton, 58 Extinction events, 301, 543 Extracellular enzymes, 355, 778 Extracellular polymeric substances (EPS), 183, 359, 406, 548, 625, 631, 656, 698, 699 calcification, 700 structure, 617 Extreme environments, 362 Extremophiles, 361, 467, 871

SUBJECT INDEX F Facultative aerobes, 9 anaerobes, 9, 460 photosynthetic bacteria, 464 Faint early sun, 829 Farbstreifensandwatt, 882 Farnesane, 171 Farrea occa, 799 Fascichnus, 121 F. frutex, 122 Fatty acids, 171, 235 Favosamaceria, 637, 642 Fe. See Iron (Fe) Feldspar, weathering, 144 Fenestral fabrics, 197, 643 Fenestral microfabrics, 672 Fermentation, 50, 232, 596 iron reduction by, 371 Ferredoxins, 559, 705 Ferribacterium limneticum, 370 Ferric iron. See Iron. Ferric oxyhydroxides (ferrihydrite). See Iron. Ferritrophicum radicicola, 368 Ferroan calcite, 721 Ferroan carbonates, 785 Ferroglobus placidus, 369 Ferroplasma acidiphilum, 857 Ferroplasma acidophilum, 362 Ferrous iron. See Iron. Fertilizer, 687 Fiber-optic sensors, 743 Fibrobacter, 90 Filaments, cyanobacteria, 307, 548 subsurface, 851 Filoreta marina, 749 Firmicutes, 81, 83, 86 Fischer–Tropsch reactions, 512, 701 FISH. See Fluorescence in situ hybridization (FISH) Fish, 767 Fisherella, 838 Fixation, 442 of geobiological specimens, 441 Flagellate, 747 Flavobacteria/Cytophaga, 84 Flavobacterium, 84 Flintstones, 273 Floccules, 787 Flood basalt volcanism, 302 cause for mass extinctions, 302 Florida Bay, 249 Florida Shelf, 249 Florideophyceae, 12 Florideophytes, 12 Flowstones, 836 Fluid chemistry, at cold seeps, 282 Fluid inclusions, 315, 771, 831 Fluidization, 663 Fluid migration, at cold seeps, 279 Fluids, 456 geothermal, 467 hydrothermal, 456 Fluorescence in situ hybridization (FISH), 146, 577, 593, 839, 373 applications in geomicrobiology, 383 Fluorescence microscopy, 374 Fluorosis, 476 Fluxes, organic carbon, 231 Foraminifera, 123, 393, 747 carbonate production, 394 denitrification, 394 endoltithic and euendoltithic, 119

endosymbionts, 394 Mg/Ca, 334 role in bioerosion, 118 Force microscopy, 772 Formaldehyde, 575 oxidation system, 576 Formate, 282, 702, 704 dehydrogenase, 575 Fossilization, of organic matter, 195 role of biofilms, 136 Fossil, Lagerstätten, 296, 785 preservation, 785 by iron sulfides, 493 in amber, 31 stromatolites, 225 Fourier transform infrared spectroscopy (FTIR), 616 Framboids iron sulfides, 488, 671 size distributions, 490 Franceville basin, natural radioactivity, 751 Francolite, 191, 733 Frankia, 86 Frasnian–Famennian boundary, 543 Freshwater and continental stromatolites, 622 Friedmanniomyces, 402, 404 Frondomorpha, 345 Frustules, diatom, 326 Frutexites, 396 F. arboriformis, 397 F. microstroma, 397 Fucoxanthin, 327 Fulgensia fulgens, 406 Fullerenes, 545 Fulvic acids, 597, 835 Fumaroles, 467, 473 Fungi, 31, 114, 348, 401, 416, 476, 812, 835 in basalt, 108 biodeterioration, 418 biogeography, 404 endoliths, 108, 348 interactions with clay, 427 mycorrhiza, 867 physiology water light temperature CO2, 405 role in biodeterioration, 114 role in bioerosion, 118 role in element cycles, 421 role in volcanic rock weathering, 145 symbioses, 428, 867 Fusarium, 420 F. oxysporum, 433 Fusulinids, 393 G Galena (PbS), 191, 857 Gallionella, 191, 272, 411, 529, 681 G. ferruginea, 104, 367, 368, 411, 588 Gallionellaceae, 411 Gamete, 533 Gammacerane, 157, 173 Gamma-proteobacteria, 83, 90, 540, 877, 879 Gamma ray burst, 303, 543 cause for mass extinctions, 302 Ganymede, Jupiter’s moon, 77 Gardening, bacterial, by foraminifera, 394 Gas chromatography-mass spectrometry (GC-MS), 148, 168 Gas hydrates, 231, 284, 577, 583 Gaskiers glaciation, 299, 887 GDGTs, 158 Geastrum, 425 Geitleria calcarea, 838

917 Gels, 812 Gemma, 215 Gemmata, 85, 155 Genes, 53 biocalcification, 53 toolkit, 55 Genomes, 439, 440 Geobacter, 370, 372, 412, 681, 780, 791 G. argillaceus, 413 G. bemidjiensis, 412, 413 G. bremensis, 413 G. chapellei, 412, 413 G. grbiciae, 413 G. humireducens, 413 G. hydrogenophilus, 413 G. lovleyi, 413 G. metallireducens, 369, 371, 372, 412, 413, 682 G. pelophilus, 413 G. pickeringii, 413 G. psychrophilus, 413 G. sulfurreducens, 97, 370, 412, 413 G. thiogenes, 413 G. uraniireducens, 413 Geochronology, 413 Geodia barretti, 841, 843 Geomycology, 416, 401 George formation, 663, 664 Geothermal fields, 447, 467, 812 Geothermobacter ehrlichii, 372 Geothermobacterium, 476 Geothrix, 84, 780 G. fermentans, 370, 372, 780 Geovibrio ferrireducens, 370 Geyserites, 448, 475, 809, 812 Geysers, 447, 467, 608, 808, 809 Giant bacteria, 879 Giant clam, 463 Giardia, 341 Girvanella, 214, 216, 639, 671 Glacial warming, Mg/Ca, 335 Glaciations, cause for mass extinctions, 302 Glaciosphere, 726 Glass, 103 volcanic, 103, 143 Glass alteration, 146 Glaucophytes, 10, 13 Globigerinoides G. bulloides, 334 G. ruber, 334 G. sacculifer, 334 Globodendrina, 124 Gloeocapsa, 348, 403 Gloeocapsomorpha prisca, 153, 157 Glucose, oxidation of, 48 Glushinskite, 424 Glutamic acid, role in biomineralization, 54 Glycerol dialkyl glycerol tetraethers (GDGTs), 158 Glycerol ether lipids, 152, 158 Glycine, 292 Glycolaldehyde, 292 Glycolysis, 596 Glycosylase, 534 Goethite, 191, 424 at hot springs, 448 Gold, 183, 426, 433, 856 Golden algae, 750 Gold labelling, 483 Gomphonema, 328 Gondwanaland, 206, 434, 543 Gongrosira, 11 Gordonophyton, 215 Graminaceous plants, iron uptake, 793

918 Granite, 548 endoliths in, 348 deep biosphere, 873 Graphite, 512 d13C-values, 512 Graphitization, 236 Grazers, bioerosion, 119 Grazing, 126, 352, 481, 782 Great oxidation event, 294, 436, 543, 688, 736 Great plate count anomaly, 90 Great Salt Lake, 766 Green algae, 10, 401 biomarkers, 157, 171 Greenalite, 92 Green and purple sulfur bacteria, 878 Greenhouse effect, role of methane, 278, 577 world, 239 Green nonsulfur bacteria, 736 Green sulfur bacteria (Chlorobiaceae), 736 biomarkers, 160, 171 Greigite, 42, 191, 486, 537, 538, 705 Groundwater, 448 Sr-isotopes, 516 Grypania, 296 G. spirali, 341 Guano, 838 Guild, 592 Gunflint BIF, 92 chert, 341 formation, 564 Gymnamoebae, 747, 748 Gymnosolen, 648 Gymnosperm, biomarkers, 173 Gypsum, 191, 523, 831, 836 crust, 348 H H2. See Hydrogen Habitat, 592 Hadean, 293 ocean, 703, period, 202 Haeckel, 702 Haemophilus influenzae, 533 Halanaerobacter, 784 Halanaerobiales, 440 Haliclona cymiformis, 13 Halide transformations, 428 Halimeda, 243, 245, 256, 589 Haliotis, 267 H. asinina, 55 Halite, 314, 437, 831 endoliths in, 350 extraterrestrial, 315 Halkieria, 61 Halley, 573 Halloysite, 424 Haloarchaea, 314, 437, 767 Haloarcula marismortui, 439 Halobacteriaceae, 364, 440, 767 Halobacteria, 314, 364, 437 Halobacterium, 314, 315, 437, 438, 439, 440 H.noricense, 314 H. salinarum, 314, 438, 873 Halococcus, 314, 438 H. dombrowskii, 314 H. salifodinae, 314, 438 Halogenated compounds, 596 Halogenated gas emission, cause for mass extinctions, 302 Halomonadaceae, 439

SUBJECT INDEX Halomonas, 438, 767 H. meridiana, 337, 678 Halophages, 84, 767 Halophiles, 159, 314, 362, 437 genome of, 439 biomarkers, 174 fungi, 767 Halorubrum, 439 Halosimplex carlsbadense, 314 Halothiobacillus, 878 Halotolerant, 767 Hangenberg event, 543 Haptophytes, 10, 15, 277 biomarkers, 173 Hardwater creeks, 24 Haslea, 153 Hautschatten, 136 HBI alkanes, 156 HCl, 830 HCO3–, 777 Heap reactors, 183 Hedstroemia, 215 Heliobacteria, 83, 86, 736 Helminthoidichnites, 886 Hematite, 92, 191 Hemiaulus, 34, 329 Hemicellulase, 357 Hemicellulose, 597 Hemichloris, 348 Heminthoidichnites, 888 Hemoproteins, 560 Heppia, 403 Heterochone calyx, 802 Heterococcus, 12 Heterocycles, in meteorites, 572 Heterocysts, 86, 307, 686 Heterokont algae, 10 Heterotrophy, 230, 716 Hexactinellid sponges, 59, 62, 797 Hexangulaconularia, 62 Hexasterophora, 797 Hg, isotope fractionation, 506 Highly branched isoprenoids (HBI), 156, 172 High-Mg calcite, 239, at cold seeps, 284 High Performance LC-MS (HPLC-MS), 169 High-temperature iron reduction, 464 nitrogen fixation, 464 Hirnantian ice age, 543, 819 Hirondella gigas, 738 Histology, 441 Holophaga, 2 H. foetida, 2 Homoacetogens, 1, 702 Hopanes, 159, 885 Hormathonema, 348 H. violaceo-nigrum, 350, 352 Hormogonia, 307 Host, symbiosis, 866 Hot spots, volcanic, 456 Hot springs. See Hydrothermal. Hot vents. See Hydrothermal. Huber–Wächtershäuser reaction, 706 Humboldtine, 424 Humic acids, 835 Humidity, 351 Humification, 835 Humus, 597 Hunsrück Slate, 785 Huntite, 262 Huronian, 815 Hyalonema, 798 H. sieboldi, 802

Hyaluronidase, 357 Hybrid crust stromatolites, 640 Hydrate Ridge, 42 Hydration of minerals, 458 Hydrocarbons biomarkers, 169 biodegradation, 236 at cold seeps, 282 Hydrocerussite, 424 Hydrogen, 50, 451, 597, 702, 781 atmospheric, 452 at cold seeps, 282 in marine hydrothermal environments, 457 metabolism, 452 oxidizers, 460 radiolytic generation, 451 role in Earth’s oxidation, 452 role in photosynthesis, 453 sources, 451 transfer, 453, 597 Hydrogenase, 559 Hydrogenobacter, 82 Hydrogen peroxide, 9 Hydrogen sulfide, 457, 477, 487, 706, 781 autooxidation, 523 at cold seeps, 282 in marine hydrothermal environments, 457 oxidation, 779 release, cause for mass extinctions, 302 Hydrolytic enzymes, 777 Hydromagnesite, 261, 424 Hydrosulfide, 704 Hydrothermal activity, 732 alteration, 469 environments, 456, 467 ancient, 454 marine, 456 terrestrial, 447, 467, 808 explosions, 473 fluids, 457 chemical composition of, 474 mounds, 704 plumes, 464 sinter, 808 springs, 224, 447, 467, 703, 808 vents, 278, 456, 751, 878, 901 fauna, ancient 455 methane oxidation at, 43 Hydroxyacids, 571 Hydroxyarchaeol, 37, 175, 885 b-Hydroxyaspartic acid, siderophores, 566 a-Hydroxycarboxylic acids, siderophores, 566, 571 Hyella, 121 H. balani, 119, 121 H. caespitosa, 121 H. gigas, 122 Hymenaea, 26, 31 Hymenelia, 403 H. coerulea, 405 H. prevostii, 349, 405 Hymenia coerulea, 349 Hymenoscyphus ericae, 425 Hyolitha, 59 Hyolithelminthes, 59 Hypersaline environments, 159, 314, 361, 437, 765 stromatolites, 622 Hyperthermophiles, 82, 364, 448, 460, 475 Hyphae, 145, 401, 416, 867 Hyphomicrobiales, 564 Hyphomicrobium, 87 Hyphomycetes, 402

SUBJECT INDEX Hypogymnia physodes, 425 Hypolithic, 349 Hysterangium crassum, 425 Hystrichospheres, 16 I Iceland, sinter 812 Ichnofossils, 481, 886 Ichnogenus, 121 Ichnoreticulina, 121 I. elegans, 119 Ichnology, 481 Ichnotaxa, 886 Ikaite, 261 Illite, 191 Immunolocalization, 482 Immunofluorescence microscopy, 483 Immunogold, 483 Immunohistochemistry, 54, 482 Immunolabeling, 482 Impacts, 69 craters, 70 cause for mass extinctions, 302 geomicrobiological influence of, 71 Inca rose, 541 Inclusions, 31 in amber, 31 oil, 885 trails, 146 Inductively coupled plasma mass spectrometry (ICP-MS), 503 Infusoria, 747 Inhibition, of carbonate precipitation, 224, 263 Inorganic carbon, 231 Insects, 33 In situ hybridization, 373 Integrated Ocean Drilling Program (IODP), 318 Interstellar medium, 292 organic matter, 572 space, 570 Interstitial waters, 742, 767 Intracytoplasmic membrane system (ICM), 575 Intraformational mud clasts, 551 Iodine radioisotopes, 280 Ion activity product (IAP), 830 Ion etching, 885 Irmicutes, 81 Iron, 367, 371, 457 biogeochemical cycle, 367 chelators, 780 cycle, Archean, 99 cycling, diagenesis, 780 Fe(III)-citrate, 780 ferric (Fe3+), geochemistry, 371 ferrous (Fe2+), geochemistry, 367, 777 as electron donor, 368 geochemistry, 370 formations, banded (BIFs), 92 hydroxide (Fe(OH)3), 191, 424, 515, 780 isotopes, in BIFs, 97 biosignatures, 504, 514 paleoredox conditions, 505 fractionation by phototrophs, 515 in marine hydrothermal environments, 457 minerals in BIFs, 92 at hot springs, 448 monosulfide, 780 ores, 92, 691 oxidation, 6, 367 assimilative, 369

by endoliths, 105 microbial, isotope fractionation, 515 phototrophic, 369 oxide dissolution, 795 formation, during weathering, 144 oxyhydroxides, 732, 777 porphyrin, 560 proteins, iron–sulfur, 559 redox couples, 368 reduction, 65, 236, 370 at high temperature, 464 role in bacterial clay authigenesis, 275 sedimentary sources, 487 shrubs, 396 siderophores, 565, 793 sorption, at bacterial surfaces, 275 sulfides formation, 486 iron-nickel sulfides, 706 paragenesis, 495 reoxidation, 496 sulfur sources, 488 textures, 488 Irpex, 420 Irrigation, 482, 782 Isochrones, 519 Isomorphic substitution, 263 Isoprenoid, 25, 235 archaeal, 158 phytane, 170 24-Isopropylcholestanes, 341, 820, 840 Isorenieratene, 151, 161, 171 Isotopes biomarkers, compound-specific, 167 biosignatures, 194 carbon, 511 effect, 579 endoliths, basalt, 107 iron, 504, 514 metals, 502 nitrogen, 513 sulfur, 513 neodymium, 517 strontium, 516 radiogenic, 516 Isotope ratio monitoring gas chromatography/ mass spectrometry (irm-GC/MS), 167 Izhella, 215 J Jarosite, 191 at hot springs, 448 Jasper, 272 Jelly rolls, 551 Jupiter, 568 K Kalicinite, 262 Kaolinite, precipitation in hot springs, 448 in shales, 787 Kathablepharids, 747 Kellwasser events, 543 and Hangenberg events, 301, 544 Kenya Rift, 812 Kerogen, 150, 169, 194, 231, 293 in cosmic matter, 293 Kerolite, 275 K-feldspars, 144 Kidney stone, 679 Kimberella, 296, 300, 886, 887 K. quadrata, 344

919 Kinetic inhibition, 460 isotope effects, 168, 512 Kinneyia, 551, 641 Kiritimati Island, 607 Klebsiella, 359 Kleptoplasts, 394 Kocuria, 529 Komokiaceans, 394 Korarchaeota, 65, 462 K/T boundary, extinction event, 302, 545 Kunmingella, 62 Kupferschiefer, 170 biomarker, 170 Kutnahorite, 262, 541 Kyrtuthrix dalmatica, 121 L Labdane, in resins, 27 Lactams, 572 Lactic acid bacteria, 597 Lactobacillus L. casei, 84 L. lactis, 371 Lactococcus lactis, 371 Lacustrine carbonates, 240 Lag deposits, of iron sulfides, 492 Lakes, 765 carbonate deposits, 240 Kivu, 827 saline, 265, 824 Tanganyika, 827 Van, 824 Vostok, 873 Lamination in black shales, 201 in shales, 786 Lampenflora, 838 Landfills, methane emissions, 583 Land plants, 300 Landsfordite, 261 Lapworthella, 61 Lasallia, 424 Last common ancestor of the metazoa, biomineralization, 55 Last common community, 711 Late Heavy Bombardment, 771 Lathamella, 62 Laurentia, 208 Lava tubes, 526 Leaching, 145, 182, 428 Lead (Pb), 448 isotopes, 518 anthropogenic, 519 dating, 518 Lecanora, 404 L. atra, 424, 425 L. muralis, 406 L. rupicola, 425 Lecanoromycetes, 403 Lecidea, 404 L. confluenta, 406 L. inops, 424, 425 L. lactea, 424, 425 L. aff. Sarcogynoides, 350, 353, 406, 407 Lectin, 484 Leguminous plant, 869 Leiolites, 617, 625 Lenticulina, 398 Lepidocrocite, at hot springs, 448 Lepraria, 404, 405 Leptospira, 84 Leptospirillum, 184, 368 L. ferrooxidans, 857

920 Leptothrix, 87, 315, 529, 535, 681 L. cholodnii, 535 L. echinata, 564 L. lopholea, 535 L. mobilis, 535 L. ochracea, 367, 368, 535 Leuconostoc mesenteroides, 84 Leucothrix, 604 Lichenothelia, 402, 404, 424 L. gigantea, 404 L. globulifera, 404 L. intermedia, 404 L. radiate, 404 Lichens, 12, 307, 348, 401, 418, 429, 867 biogeography, 404 distribution patterns, 404 morphology, 402 physiology water light temperature CO2, 405 Lichinella, 403, 405 Life, astrobiology, search for, 75 definition, 75 environmental limits, 74 origin, 701 Light, 350, 405 intensity, in rocks, 350 Lignin, 597 degradation of, 356, 420 Ligninase, 357 Lime mud, 663, 665 mound, 667 Limestone, 785 Limiting nutrient, phosphorus, 732 Limnoriidae, 902 Lingula, 496 Linguliformea, 59 Lipase, 357 Lipids, 168, 235, 701 biomarkers, 147 degradation of, 356 marine, degradation, 235 Liquid chromatography (LC), 169 Lithistida, 797 Lithocodium, 644 Lithoglypha, 406 Lithoherms, 668 Lithothamnion sp., 141 Lithotrophic, 50 Lithozoa, 747 Lixiviants, 183 Lobopodians, 59 Logatchev vent field, 461 Long-chain alkenones, 173, 278 Lophelia, 253 Lophotrochozoa, biomineralization, 56 Lost City hydrothermal field, 456, 703 Lottia scutum, 55 Low–Mg calcite, 263 at cold seeps, 284 Low-temperature fluids, 458 Low-temperature hydrothermal ore deposits, 694 Lublinite, 666 Lubomirskia baicalensis, 800 LUCA, 293 Lueckisporites, 314 Luvisols, 835 Lyngbya, 217, 308 L. aestuarii, 548 Lytoceratina, 545 M Mackinawite, 191, 486, 538, 705 Macrobioerosion, 117 Macroborers, 118 Macrocyclic alkanes, 171

SUBJECT INDEX Macrofauna, 782, 783 Macromolecular materials, 572 Macromolecules, role in biocalcification, 54 Macromonas bipunctata, 667 Macronutrients, 598 Madrepora, 253 Magmatism, 206 Magnesian calcite, 721 Magnesite, 191 Magnesium (Mg), calcite, 263 distribution coefficient, 264 in marine hydrothermal environments, 457 in seawater, 331 in carbonates, 263 mid-ocean ridge, 332 Magnetic field, 538 Magnetic force microscopy (MFM), 775 Magnetite, 92, 191, 537, 538, 682 as biosignature, 192 Magnetobacterium bavaricum, 539 Magnetobacterium gryphiswaldense, 539 Magnetobacterium magnetotacticum, 539 Magnetofossils, 540 Magnetosomes, 537 Magnetospirillum, 539 Magnetotactic bacteria (MTB), 537 Magnetotaxis, 537 Makganyene glaciation, 817 Malachite, 262 Maleimides, 152, 170 MamJ protein, 539 Mammoth Cave, 522 Manchuriophycus, 549 Manganese (Mn), 370, 424, 448, 457 carbonates, 541, 780 cycling, diagenesis, 743, 779 in marine hydrothermal environments, 457 nodules, 394 oxyhydroxide, 194 oxidation, 779 reduction, 49, 236, 779 sedimentary, 541 sulfides, 541 Manganoan calcite, 789 Mannan degradation of, 356 Mantle plumes, 207 Marcasite, 486, 494, 495 Marchandia magnifica, 33 Mariana Trench, 738 Marichromatium purpuratum, 602 Marine carbonate saturation, geological record, 217 Marine snow, 89, 258, 361 Marinoan glaciation, 888 Marinobactins, 565 Mariprofundus ferrooxidans, 105 Marls, 245, 785 Mars, astrobiology, 73, 74, 76, 146 meteorite ALH84001, 514, 540 Mass dependent sulfur isotope fractionation, 295 Mass extinctions, 301, 543 Massive sulfide deposits, 458 Mastigocoleus testarum, 119, 121 Mastogloia, 329 Mat chips, 551 Mat-related sedimentary structures, 547 Matrix-mediated mineralization, 589 Maturation, amber, 27 Mawsonites pleiomorphus, 346 Meishucunian, 63

Membrane, 708 archaeal, 65 bacterial, 82 Mesoclots, 624 Mesophiles, 364, 460 Mesostigma, 12 Messel oil shale, 785 Messenger RNA, origin, 709 Messinian salinity crisis, biomarkers, 174 Metabolism, 456 aerobic, 8 anaerobic, 48 Metagenesis, 150, 230 Metagenomics, 553 Metal, 427 acquisition, bacterial, 565 binding, microbial, 654 enzymes, 558, 711 immobilization, 428 immobilization by fungi, 428 ion enrichment, in burrows, 482 isotopes, 502 mobilization, by fungi, 428 proteins, 558 sequestration, microbial surfaces, 190 sulfide catalysts, 711 sulfide minerals, 856 transformations, by fungi, 427 Metallogenium, 563, 853 M. personatum, 564 M. symbioticum, 564 Metalosphaera, 184 Metaproteomics, 553 Metatranscriptomics, 553 Metazoa, eggs, 877 calcification, 53 silification, sponges, 796 skeletons, 58, 796 origin, 716 Meteorites, 568, 701 Methane, abiogenic, 283 activation, 37 anaerobic oxidation of, 36 aerobic (bacterial) oxidation of, 283, 575 biogenic, 66, 232, 282, 576, 702, 781 biomarkers (compound-specific isotopes) of, 175 (seep-related) carbonates, 66, 286 isotopes, 286 cold seeps, 278 monooxygenase (MMO), 575 origin, 578 release, 578 cause for mass extinctions, 302 thermogenic, 282 thiol, 704 on Titan, 77 Methanobacterium, 337 M. thermoautotrophicum, 64 Methanocaldococcus, 462 M. janaschii, 197 Methanococcoides, 37 Methanococcus, 319, 365 Methanogenesis, 65, 282, 576, 702, 711 biomarkers (compound-specific isotopes) of, 175 diagenesis, 781 reverse, 36 role in dolomite formation, 338 role in organic matter degradation, 235 reactions, 66 substrates, 282, 578

SUBJECT INDEX Methanogenium, 365 Methanol, 282, 575, 704 dehydrogenase, 575 Methanomicrobiales, 37 Methanopyrus, 448 M. kandleri, 65, 75, 364 Methanosaeta, 337, 579 Methanosarcina, 579 M. acetivorans, 2 Methanosarcinales, 37, 463 Methanothermus, 364, 476 Methanotrophs archaea, 36, 159 bacteria, 575 biomarkers, 160, 175 Methanotrophs type I, 575 Methanotrophs type II, 575 Methanotrophs type X, 576 Methylamine, 282 Methyl-coenzyme M reductase, 560, 684 Methylhopanes 2a-, 96, 97, 160 3b-, 160, 175 Methylobacter, 463, 576 Methylocaldum, 576 Methylococcus, 576 Methylocystis, 576 Methylomicrobium, 576 Methylomonas, 463, 576 Methylophaga, 463 Methylosinus, 576 Methylosphaera, 576 Methylotrophs, 575 Methyl sulfide, 706 Methyl thioacetate, 706 Methyl thiolate, 702 Mg. See Magnesium. Mg/Ca paleothermometry, 334 Mg/Ca ratios, 239, 448 Mica, 786 Micrite, 219, 664, 672, 785 auto-, 258 encrustations, 128 Microaerophiles, 460, 537 Microautoradiography, 485, 594 Microbacterium flavescens, 795 Microbial biofilms, 134, 606 biomineralization, 586 biosignatures, 189 buildups, 617, 635 calcite precipitation, 211, 223 community structure, 592 consortia in biodeterioration, 112 corrosion biosignatures, 192 decay profiles, 788 degradation, 596 diversity, 592 dolomite, 336 ecology, of caves, 599 fabrics, in rocks, 196 farming, 843 fossils, 810 laminae, 787 laminites, 817 loop, 748 mats, 134, 547, 606, 699, 733, 880 metal binding, 654 ore precipitation and oxidation, 691 population, 592 processes in biodeterioration, 112 sand chips, 551 secretions, 359 silification, 608

surface reactivity, 614 weathering, 144 Microbialites, 53, 617, 635, 698, 699, 830 Microbially induced early diagenetic cements, 785 Microbially induced sedimentary structures (MISS), 196, 547, 617 Microbiocorrosion, 657 Microbioerosion, 117 Microborers, 118 Micrococcus sp., 187, 359 Microcodium, 242 Microcoleus, 85 M. chthonoplastes, 550 Microdictyon, 61 Microfossils, earliest, 195 Microsensors, 658 Microspar, 662, 664 Microtome, 443 Mid-ocean ridges (MOR), 456 Migration, 281 Millepora, 245 Millerite, 191 Mineral, 235, 422, 733 biooxidation, 182, 691, 856 deposits, at deep sea hot vents, 456 precipitates, microbial induced, 586, 191 sorption of organic matter, 235 transformations by fungi, 422 Mineralization, 449, 596 divalent cations, 335 microbial, 274, 586, 691 Mineralized microbes, 449 Mining, burrowing organisms, 481 Minoan eruption, 825 Mississippi-Valley-Type (MVT), 693 Mitochondria, 8 Mixotrophy, in algae, 14 Mn. See Manganese. Mobergellans, 59 Mofettes, 473 Moganite, 812 Molar-tooth structure, 662 Molecular clock, diatom, 329 Molecular fossils, 147, 441 Molecular phylogeny, diatom, 329 Mollusks, 54, 59, 287 biomineralization, 54 at cold seeps, 287 Molybdenite, 857 Molybdenum, 562, 704 isotopes fractionation, 505 paleoredox conditions, 505 Molybdopterin (sulfite oxidase), 559 Mondmilch, 625, 666 Mono Lake, 363 Monomethylalkanes (MMAs), 171 Monooxygenases, 598 Monorhaphis chuni, 802 Monoterpenes, in amber, 26 Montastraea, 245 Montastrea, 129 Monteville formation, 665 Montmorillonite, 277, 424 Moolooite, 424 Moonmilk, 625, 666, 838 Moorella, 2 M. perchloratireducens, 3 M. thermoacetica, 1 Moritella, 739 M. japonica, 741 Morphology, 402 Mössbauer spectroscopy, 558

921 Mosses, 838 Motility, 326, 481 diatoms, 326 Mounds, 667, 809, 893 Mounting, of geobiological specimens, 448 Mucor, 420, 424, 425 Mucus, 481 Mud, 785 diapirs, 282 mounds, 254, 667, 893 volcanoes, 42, 279, 282, 319 Mudpots, 473 Mudstone, 785 Multicellularity, triggered by Ca, 830 Multicellular magnetotactic prokaryote (MMP), 539 Multicollector inductively coupled plasma mass spectrometry (MC-ICP-MS), 415 Murchison meteorite, 569 Murein, 82 Mussel shrimp, 767 Mutualism, 675, 866 Mycobacterium, 151, 350 Mycobiont, 401, 402 Mycogenic minerals, 424 Mycoplasma, 86 Mycoplasmatales, 564 Mycorrhiza, 416, 418, 429, 675 Mycorrhizae, 867 Mytilid mussels, 463 Myxobacteria, 87 Myxococcus xanthus, 187, 188 Myxosarcina, 307, 308 N N. See Nitrogen NAD, 575 NADH, 48, 575 NADH2, 8 Nahcolite, 262 Namacalathus, 59, 61, 296 Namapoikia, 58, 61, 296 Nama-type biota, 345 Nanoarchaeota, 65, 462 Nanoarchaeum equitans, 679 Nanobacteria, 225, 677, 695 Nanobes, 677 Nanocrystals, 537, 540, 681 Nanoorganisms, 677 NanoSIMS, 594 Nanospheres, 784 Nanovesicles, 677 NASA, 74 Natrialba, 438 Natron, 262, 824 Natroniella, 2 Natronincola, 2 Natronococcus, 65, 438 Natronomonas, 438 N. pharaonis, 439 Natural attenuation, 598 Natural fission reactors, 751 Natural radioactivity, 751 Nautilus, 258 Navicula, 153, 328 N. veneta, 328 Near-field scanning optical microscopes (NSOMs), 776 Needle fiber calcite, 722 Nemiana simplex, 344 Nemrut, 826 Nenoxites, 887 Neodymium (Nd), isotopes, 517 Neoichnology, 481

922 Neopelta, 799 Neoproterozoic, 206 salt deposits, 771 trace fossils, 886 Nepheloid flows, 787 Nesquehonite, 261 Neurospora sitophila, 433 Neutralization, of acid rock drainage, 7 Neutrophiles, 460 iron oxidizing, 368 iron reducing, 372 Nickel, 561, 684 famine, 296 removal from acid mine drainage, 7 Nickel-containing hydrogenases, 685 Nicotinamide dinucleotide (NAD), 702 NiFe hydrogenase, 559, 560 Ni-protein I, 37 Nitobacter, 83 Nitrate (NO3–), 462, 686, 777 ammonification, 779 reduction, 687 reduction, microbial, by Fe(II) oxidation, 368 Nitrification, 513, 685, 687, 779 nitrogen isotope fractionation, 513 Nitrifiying bacteria, 87 Nitrile hydratase, 559 Nitrite (NO2–), 43, 686, 777 oxidizers, 779 Nitrobacter, 114, 186, 529, 687, 779 Nitrococcus, 687 Nitrocystis, 687 Nitrogen, 230, 686 cycling, diagenesis, 779 fixation, 307, 513, 686 in cyanobacteria, 307 high temperature, 464 nitrogen isotope fractionation, 513 gas (N2), 686, 779 heterocycles, 572 isotopes (d15N), 513 oxides (N2O, NOx), 686 Nitrogenase, 307, 686 H2-production, 453 Nitrosomonas sp., 186, 529, 687 N. europaea, 83 Nitrospina, 687 Nitrospira, 539, 687 Nitzschia, 329 Nonenzymatic carbonate deposits (mud mounds), 674 Nonprotein amino acids, 570 Nostoc, 308, 401, 838 Nostocales, 310 Nostochopsis lobatus, 350 Nuclear magnetic resonance (NMR) spectroscopy, 360 Nucleation, carbonates, 224 Nucleic acids, 235, 709 DAPI stain, 146 Nuclides, radiogenic, 516 O O. See Oxygen. Obruchevella, 215 Obsidian, 144 weathering, 144 Ocean, 25, 769 acidification, 54, 129, 141 alkalinity, ancient, 23, 829 Ocean Drilling Program (ODP), 318, 733 Oceanic anoxic events (OAEs), 159, 203, 733 biomarkers, 174 Ocean water chemistry, ancient, 300, 769, 829

SUBJECT INDEX Ochrobactins, 565 Ochrolechia parella, 425 Oikomonas, 748 Oil inclusions, 885 shales, 202 spills, 598 Okenane, 151, 161 Okenone, 151, 172 Oleanane, 158 Oligonucleotide probes, 374, 378, 577 sequence, 555 Oligotrophic, 733 Olivines, 145, 457, 702 in marine hydrothermal environments, 457 volcanic rock weathering, 145 Oncoids, 212, 809 Oocardium, 11 Oogamy, diatoms, 327 Ooids, 53, 225, problem 252 Ooze, 258 Opal, 60, 191, 448, 608, 812 Opaline silica, 812 Open marine stromatolites, 617, 635 Optodes, 743 Orbulina universa, 334 Ordovician, 733 Oreaster, 245 Organic acids, weathering, 144 Organic carbon. See organic matter. Organic matter (OM), 149, 698, 700 anaerobic degradation of, 48 cycling, 230 decomposition 234, 420, 777 role in carbonate diagenesis, 140 diagenesis, 149, 777 macromolecules, 54 preservation, 232 respiration, 8 Organic microanalysis, 884 Organic molecules biomarkers, 147, 167 biosignatures, 189 in cosmic matter, 292, 568 origin of life, 701 Organofilms, 135, 697 Organohalogens, 598 Organometals, 420 transformations, 427 Organomineralization, 53, 698, 837 vs. biomineralization, 698 Organotrophs, 122 Orgueil carbonaceous chondrite, 573 Origin of Life, 701 RNA world, 763, 709 role of hydrogen, 452 thioester world, 706, 876 Orion cloud, 293 Orthogonum, 124 Ortonella, 215 Oscillatoria, 83, 476, 624, 812 O. spongeliae, 842 Oscillatoriales, 310 Osedax, 901 Osmoregulation, 767 Osmotic pressure, 364 Ostreobium, 121, 125 O. quekettii, 119, 121, 122, 123, 128, 129 Ostwald ripening, 812 Oxalates, 426 Oxalic acids, 144 Oxalotrophic pathway, 723

Oxic, 9 zone, 778 Oxidative stress, strategies used by acetogens, 3 Oxobacter, 2 Oxycline, 576 Oxygen, 230, 282 aerobic respiration, 8 crisis, 436 depletion, black shales, 203 detoxification, 9 diagenesis, 777 exposure time, 236 great oxidation event, 294 rise in Ediacaran time, 346 Oxygenic photosynthesis, 702, 736 early, 96 model, 95 Oxyhemocyanin, 559 P Paenibacillus polymyxa, 371 Palaeobolus, 62 Palaeoconchocelis starmachii, 125 Palaeopascichnida, 343 Palaeopascichnus delicatus, 344 Palaeopasichnus, 887 Palaeoscolecida, 59 Palaeozygnema spiralis, 32 Palagonite, 103 Paleobathymetry, bioerosion indicator, 121 Paleocene/Eocene thermal maximum (PETM), 159, 287 Paleophragmodictya, 61 Paleoproterozoic, 161, 296, 831 Paleosinters, 813 Paleosolex, 298 Paleothermometry, seawater, 334 Sr-isotopes, 517 Palimpsest ripples, 548 Pangea, 545, 771 Pantetheine, 876 Pantoea agglomerans, 371 Paracarinachitids, 59 Paracercomonas, 749 Paracoccus, 687 P. denitrificans, 8 Paragenesis, of iron sulfides, 495 Parasites, 746 Parasitism, 866 Parmelia, 404, 424 P. conspersa, 424, 425 P. subrudecta, 425 P. tiliacea, 424 Parmotrema, 404 Particulate methane monooxygenase (pMMO), 575 Particulate organic carbon (POC), 778 Particulate organic matter (POM), 235, 317 Passive margin, 206 Pavona clavus, 517 Paxillus involutus, 429 PCO2, 829 PCR (polymerase chain reaction), 711 P/C ratios, 732 Pearceite, 191 Peccania, 403 Pecten, 267 P. maximus, 120 Pectin, 597 degradation of, 356 Pectinase, 357 Pedogenesis, 833 Pedogenic carbonates, 721

SUBJECT INDEX Pedosphere, 833 Pelagophytes, 156 Pelletal fabrics, 787 Peloids, 196, 225, 669, 672 Peltigera rufescens, 405 Peltula, 403 Penicillium, 420, 424, 425 P. coryliphilum, 424, 425 P. frequentans, 424 P. simplicissimum, 424, 425, 426 P. simplicissimum Verrucaria, 424 P. steckii, 424 Penicillus, 243, 245 Pennate diatoms, 326, 881 Pentamethylicosane (PMI), 37, 159 Peptides, 707 bonds, 829 synthesis, 707 Peregrinella, 256 Peridinin, 16 Peridotite, 456 Periplasmic space, 82 Perlucin, 54 Permafrost, 726 Permian, 733 salt deposits, deep biosphere, 313 Permian–Triassic extinction, 301, 544 Perpetual spouters, 447 Pertusaria corallina, 424, 425 Petalonamae, 345 Petee ridges, 549 Petrobactin, 567 Petroleum, 148 degradation, 151 formation, 148 pH, 141, 448 role in calcium carbonate precipitation, 223 Phaeococcomyces, 402 Phaeophila, 121 Phaeophyceae, 14, 747 Phaeosclera, 402 Phagotrophy, of dinoflagellates, 15 Phanerochaete, 420 P. chrysosporium, 420 Phanerozoic, 125 mass extinctions, 301, 543 shales, 788 Phase detection microscopy (PDM), 775 Phellinus, 428 Phenols, 598 Phoma, 402 glomerata, 424 Phormidium, 85, 217, 308, 438, 476, 477, 612, 624, 812 Phosphate, 191, 707, 732, 780, 877 coatings, 782 concretions, 788 early animal skeletons, 60 minerals, 733 Phosphatocopida, 59 Phosphogenesis, 732 Phosphoglycerate (PGA), 512 Phosphorites, 296, 732 Phosphorus (P), 732, 780, 788 Phosphorylation, 707 Photic zone euxinia (PZE), 151, 172 Photobacterium, 739 Photobionts, 12, 401, 402 Photochemical methane oxidation, 577 Photoorganotrophs, 738 Photosynthesis, 231, 405, 736 role in BIF deposition, 95 role in Earth’s oxygenation, 294 Photosystem, 86, 702

Photothermal microspectroscopy (PTMS), 775 Phototrophs, 113 bacteria, in deep sea hydrothermal environments, 464 endoliths, 121 Fe(II) oxidation, 780 role in biodeterioration, 113 anaerobic, 51 PHREEQE, 827 Phycobilins, 12, 307 Phyllisciella, 403 Phylloceratina, 545 Phylogenetic tree, 81 Phyrococcus, 319 Physcia adscendens, 402 Phytane, 170 Phytophtora, 120 Phytoplankton, 235 biomarkers (compound-specific isotopes) of, 170 pico-, 309 Phytosiderophores, 793 Phytozoa, 747 Picrophilus, 362 Piezophilic bacteria, 364, 738 Pigments, 235, 307, 327 in cyanobacteria, 307 in diatoms, 327 Pilatus mountain, 667 Pilbara, 195 Pillow lavas, 146 Pimarane, in resins, 27 Pisoids, 809 Pit marks, 106 in basalt (glass), 106 Plagioclase, in marine hydrothermal environments, 457 Planctobacteria, 90 Planctomyces, 85 Planets, 293 astrobiology, 75 Plankton, 235 Plankton-derived organic matter, 235, 778 Planolites, 258 Planothidium frequentissimum, 328 Plant, 10, 144, 157 litter, 597 nutrition, 833 rise of land plants, 300 role in weathering, 144 roots, 834 terrestrial, biomarkers, 173 Plasmodium, 746, 747 Plasticity, 785 Plastids, 327 Plate counts, 592 Platforms, carbonate, 243 Platygyra, 245 Platysolenites, 60 Playas, 765 Plectonema, 217 P. boryanum, 433 P. terebrans, 121, 122 Plectonema (Leptolyngbya) terebrans, 122 Pleosporales, 402 Pleurocapsa, 476 Pleurocapsales, 310 Pleurosigma, 153 Pleurotus, 420 Plutonium, 795 PMI (pentamethylicosane), 37, 159 Pocillopora, 245 Pock marks, 279 Podzols, 835

923 Poikiloaerobic, 203 Poikilohydric, 405 Polar climate, 364, 726 endoliths in, 348 Poly-beta-hydroxybutyrate (PHB), in Beggiatoa, 111 Polychaete worm, 463 Polyclonal, antibodies, 483 Polycyclic aromatic hydrocarbons (PAHs), 420 in cosmic matter, 292 in meteorites, 572 Polygalacturonans, 597 Polyhydrate calcium oxalate, 723 Polyhydroxylated compounds, in meteorites, 571 Polymerase chain reaction (PCR), 315, 577, 711 Polymerization, 812 amber, 26 Polymuds, 896 Polyols, 292, 571 Polyphosphate, 877 in Beggiatoa, 111 Polyplacophorans, 902 Polysaccharides, 235 Polysulfide, 791 Polyunsaturated fatty acids, in piezophiles, 740 Pore sizes, 742, 834 Pore water, 139, 742 migration, at cold seeps, 279 zonation, 744 Porifera, 796, 840 Porites, 120, 245 P. lobata, 123 P. lutea, 123 Porphyra, 12 Porphyrin, 148, 559 Poseidon, at Lost City, 459 Posidonia, 243 Posidonia shale, 785 Prebiotic chemistry, 701 Precambrian, 150, 160, 450, 809 boundary, 733 critical intervals, 293 rocks, 150 Precambrian–Cambrian boundary, 733 Precipitation, authigenic, clay, 274 biominerals 411, 586, animals, 53, 58, 796 carbonates, 223, 238, 262 cyanobacterial, 211 dolomite, 336 mediated by EPS, 360 pedogenic, 721 fungal, 416 gold, 433 methane-derived, 278 microbialites, 617, 635 nanocrystals, 681 iron oxides, 511, 537 on microbial surfaces, 614 mud mounds, 667, 893 ores, 691 iron sulfides 486 phosphorite, 732 sinter, 608, 808 tufa, 889 Predation, 481 role as evolutionary driving force, 54 role in bioerosion, 125 Preservation, 136, 189 of microbes in sinter, 448 of microbial structures, 194

924 Preservation (Continued ) of organic matter, 202 of organisms in resin, 31 soft tissue, 493 Priapulida, 59 Primaric acid, in resins, 26 Primary production, 231 Primordial acids, 829 ocean, 830 organic soup, 876 Pristane, 171 ProbeBase, 380, 593 Prokaryotes archaea, 64 bacteria, 81 Propane, 282 Propionibacterium pentosaceum, 84 Propionyl CoA, 598 Prosthecobacter, 85 Protease, 357 Protection bioprotection, 185 of mud surfaces, 787 of organic matter (OM), 232 Proteins, 235 degradation of, 356, 597 Proteobacteria, 86, 576 in marine hydrothermal environments, 461 Proterozoic, 124, 202 Protista, 746 Protium, 168 Protoacetogens, 712 Proto-archaea, 711 Proto-bacteria, 711 Protoctista, 747 Protoenzymes, 704, 708 Protohertzina, 61 Protomethanogens, 712 Protonation, 616 Proton motive force, 596, 707 Protophyta, 746 Prototheca, 10 Protozoa, 746 Proustite, 191 Pseudallescheria boydii, 425 Pseudanabaena, 308 Pseudendocloniopsis, 10, 11 Pseudendoclonium, 10, 11 Pseudoalterobactins, 566 Pseudoinclusions, in resins/amber, 29 Pseudo-kutnahorites, 541 Pseudomonas, 81, 187, 315, 687 P. aeruginosa, 433 P. bipunctata, 667 P. carboxdovorans, 83 P. denitrificans, 83 P. maltophilia, 433 P. stutzeri, 186 P. GS–15, 83 Psychromonas, 739 Psychrophiles (cryophiles), 364, 460 Pt-group elements, 546 Pulsating springs, 447 Purcell Supergroup, 663 Purple bacteria, 736 sulfur bacteria, biomarkers, 161, 172 Push–pull tests, 576 Pyrenopsis, 402 Pyrite, 191, 202, 488, 704, 750, 856 aggregates and clusters, 491 in black shales, 202 at cold seeps, 284

SUBJECT INDEX concretions and pyritic layers, 491 framboids, 488, 788 oxidation of, 6, 856 polyframboids, 491 Pyrite-Pulled Theory, 706 Pyrobaculum, 448 P. islandicum, 370 Pyrococcus, 448, 476 P. abyssi, 195, 197 P. furiosus, 370, 433 Pyrodictium, 448, 462 Pyrolobus, 462, 476 Pyrophosphate, 703, 706 Pyrosequencing, 593 Pyroxenes, 145 in marine hydrothermal environments, 457 volcanic rock weathering, 145 Pyruvate, 596 Q Qiongzhusian (Atdabanian), 63 Quantum dots (qdots), 485 Quartz, 786, 812 rhinds, 353 R Racemic mixtures, 75 Radioactivity, 751 sulfur isotopes, 864 waste disposal, 423 resistance, 365 Radiocarbon dating, 348, 415 Radiogenic isotopes, 516, 751 Radiolaria, 747, 754, 788, 797 Radiolarite, 272, 754 Radiometric age determination, 415, 516 Radionuclide, 427 Radioresistance, 365 Ralstonia, 81 R. eutropha, 83 Ramalina maciformis, 405 Raman effect, 755 shift, 776 microscopy, 194, 754 Rambergite, 541 Ramps, carbonate deposits, 247 Rangea schneiderhoehni, 344 Rangeomorpha, 345 Raphid diatoms, motility, 326 Raphinesquiiana alternata, 125 Rayleigh scattering, 755 criterion, 757 Razumovskia, 215 Reaction rates, 744 Reactive barriers, 7 Reactivity of crystalline phases, 780 of organic matter, 781 Recalcitrant substrates, 835 Red algae, 12 Redfield-Richards ratio, 139, 686 Redox chain, 49 couples, iron, 368 potentials, 49 aerobic respiration, 8 reactions, 460 microbial, 461 zonation, 743 Red tides, 734

Reduction spheroids, 761 Reefs, 225, 249, 257, 733, 762 carbonate budget, 130 early Cambrian, 218 Remineralization, of organic matter, 232, 234, 596 during diagenesis, 777 Renalcis, 215, 399, 643, 671 Replication, early life, 709 Reservoirs, of organic carbon, 230 Residence time, 732 Resin, 25 maturation, 27 Resinite, 24 Resonance Raman, 558 Respiration, 8, 234, 406, 596, 777 Respiratory chain, 8, 596 Resting cysts, of dinoflagellates, 16 Resting stages, algae, 14 Restoration agents, 115 Reversed tricarboxylic acid cycle, 171 Rhacodium, 402 Rheology, 663 Rhizaria, 10, 393, 747 Rhizobia, 87 Rhizobium, 686, 869 Rhizocarpon, 424 Rhizomorphs, 723 Rhizon samplers, 742 Rhizopoda, 747 Rhizopogon rubescens, 424 Rhizopus, 359, 424 Rhizosolenia, 153, 157, 329 Rhizosphere, 144, 577, 834 Rhodobacter, 81, 369 Rhodochrosite (MnCO3), 541, 780, 817 Rhodomicrobium, 87 Rhodophyta, 12, 763 Rhodopseudomonas, 153, 687 R. palustris, 97 Rhodospirillum, 83 R. rubrum, 561 Rhodovibrio, 602 Rhodovulum robiginosum, 369 Rhombocorniculum, 61 Rhopalia, 121 Rhynchonelliformea, 59 Rhynie Chert, 273, 300 Ribosomal gene, 376 Ribosome, 709 Ribulose biphosphate carboxylase/oxygenase (“Rubisco”), 307, 512, 737 Ribulose monophosphate pathway, 575 Rice paddies, 576 Richelia, 329 Riftia, 463, 879 R. pachyptila, 869 Rinodina, 404 Rivularia, 671 RNA world, 709, 763 Rodinia, 206, 300, 771 Roll-up structures, 550, 817 Root exudates, 833 Rootsicles, 838 Rosalina, 124 Roscoelite, 761 Roseobacter, 81, 83 Rossella R. fibulata, 802 R. nuda, 802 Rubisco, 307, 512, 737 Ruminococcus, 2 Rundkarren, 242 Runzelmarken, 641

SUBJECT INDEX S S. See Sulfur Sabkha, 882 Saccharomyces, 69, 359 S. cerevisiae, 416 Saccomorpha S. terminalis, 120 Sagittarius B cloud, 292 Saline brines, Sr-isotopes, 517 carbonate deposits, 241 environments, 362 microorganisms, 437 lakes, 765 Salinibacter, 440 S. ruber, 438, 440 Salinity, ocean, 769 Salmonella, 81 S. enterica, 534 S. typhimurium, 533 Salt, 765, 768 deposits, microorganisms in, 313 Salterella, 60 Salt lakes, 24, 765, 824 Saltpeter, 688 Sample preparation, for microscopy, 441 Sand, 833 clasts, 551 Sandberg-Cycles, 239 Sandstone, 348 Sarcinomyces, 402 Sarcogyne cf. austroafricana, 406 S. pruinosa, 350 Saturation index (SI), 23, 830 Saturation state, 781 of CaCO3, 23 carbonate minerals, 141 Scanning electron microscopy, 348 Scanning force microscopy, 772 Scanning near-field acoustic microscopy (SNAM), 775 Scanning probe microscopy, 772 Scanning thermal microscopy (SThM), 775 Scanning tunneling microscope (STM), 772–773 Scavenging, 481 Scenedesmus, 10, 12 Schizosaccharomyces pombe, 682 Schizothrix, 214, 243, 620 S. arenaria, 438 Schwertmannite, 191 Sciadopityaceae, 31 Sclerochronology, 252, 414 Scolecobasidium, 402 Scolymastra joubini, 802 Scoponodus, 61 Scytonema, 243, 644 S. julianum, 838 Sea level, 129 changes, cause for mass extinctions, 302 Sea surface temperature (SST), records of, 153, 334 Sea urchins, biomineralization, 54 Seawater, 20 Sr isotopes, 517 Secondary endosymbiosis, diatom, 329 Secondary ion mass spectrometry (SIMS), 594, 883 Secondary metabolites, 597 Secondary minerals, 144 formation, 777 precipitation, 782, 783 Sectioning, of geobiological specimens, 443 Sediments, 128, 231, 236 carbonates, 224, 238 cohesiveness, 782

diagenesis, 778 marine, organic carbon degradation, 235 organic carbon, 779 organic matter (OM), 698, 699 rocks, organic carbon degradation, 236 Sediment-water interface, 782 Seep carbonates, 286 fauna, 285 microorganisms, 36 Selective preservation, 235 Selenium, 426, 784 Selenocysteine, 785 Semidendrina, 124 Sepia, 258 Sequestration, of cations in EPS, 360 Serine pathway, 575 Serpentinization, 279, 282, 451, 458 Serpula himantioides, 424 Sesquiterpenes, in amber, 26 Shaking, seismic, of sediments, 666 Shales, 785 black shales, 201 Shamovella/Tubiphytes, 644 Shark Bay, 243, 617, 622 Sheaths, cyanobacterial, calcification of, 212 Shelves, carbonate, 243 Shewanella, 105, 370, 372, 514, 681, 739, 791 S. frigidimarina, 105 S. oneidensis, 370 S. putrefaciens, 682 Shores, carbonate deposits, 241 Shortite, 262 Shrinkage structures, of sediments, 666 Shuguria, 215 Siberian traps, 545 Siboglinid tube worms, 902 Siderastrea, 243 Siderite (FeCO3), 721, 780, 792 Siderophores, 145, 565, 793 Sideroxydans paludicola, 368 Silica, 326 aluminosilicate, clay biomineralization, 276 biomineralization, sponges, 796 in Archean ocean, 94 in marine hydrothermal environments, 457 minerals, at hot springs, 448, 608, 808 residue, 470 Silicatein, 800 Silicate weathering, 143, 829 Silicic acid, 801 Siliciclastic biolaminites, 552 Siliciclastic stromatolites, 641 Siliciclasts, 202 Silicification, of microbes, 192, 450, 608, 812 Silicispongea, 797 Silicoflagellates, 14, 797, 808 Silicon, 797 Silled basins, 203 Silt, 833 Silver, 426, 856 SIMS (secondary ion mass spectrometry), 883, 594 Sinkholes, 521 Sinosachites, 61 Sinotubulites, 58, 61 Sinters, 447, 467, 469, 808 as geyser caprock, 447 at hot springs, 448 SiO2, 829 Siphogonuchites, 61 Siphonophycus, 217 Skeletons, 224 early animals, 58

925 evolution, 53 of early animals, mineralogy, 59 of early animals, morphology, 61 genetic toolkit, 56 Skeneimorphs, 902 Slime, 359 Slope, carbonate, 256 Sloughing, of biofilms, 325 Small shelly fauna, 814 Small shelly fossils (SSFs), 58, 814 Small subunit ribosomal RNA, 593 Smectites, 191, 274 microbial formation of, 145 SMTZ, 42 Snowball Earth, 94, 286, 296, 814 Soda, 824 lakes, 21, 224, 362, 765, 824, 829 ocean, 23, 224, 300, 829 Soft tissues, 785 fossilization, 136 preservation, 493 Soil, 234, 732, 833 aggregation, 427 formation, 597 formation in Ediacaran time, 346 genesis, 597 microbiology, 835 mineralogy, 833 neogenesis, 145 organic matter, 234 profiles, 834 properties, 427 role of fungi in, 422 type, 834 Solar system astrobiology, 75 habitable zone, 75 Solenastrea, 243 Solentia, 620 S. achromatica, 119 Solfataras, 473 Sols, 812 Solubility, 262 carbonates, 262 product, 830 Soluble methane monooxygenase (sMMO), 575 Sound, 253 Source rocks, black shales, 201 Spaerotilus, 87 Sparry crust, 639 Sparus aurata, 768 Spatial/seasonal oscillations, 51 Speleogenesis, 521 Speleothems, 529, 636, 666, 809, 836, 889 Spelter, 905 Sphaerozoum punctatum, 334 Sphalerite (ZnS), 191, 458, 695, 857, 905 Spheroliths of calcite, 724 Spicopal, 798 Spiculites, 273 Spirilla, 538 Spirochaeta thermophila, 85 Spirochete, 84, 85 Spiroplasma, 86 Spirula, 258 Spirulina, 309, 438, 476, 812 Sponges, 158, 797, 840 bioerosion, 124 biomarkers , 158 biomineralization, 796 microbes, 840 spicules, 788, 796 Spongiostrome, 639

926 Sporobolomyces salmonicolor, 433 Sporomusa, 2 Sporosarcina pasteurii, 187 Springs, carbonate deposits, 240 hot, 447, 467, 808 intermittent, 447 tufa, 889 Squalane, 158 Stable isotope probing (SIP), 594 Stable isotopes, 167, 264 Stable sulfur isotopes, 864 Stagnation, black shales, 203 Staining, 441 of geobiological specimens, 444 Stalactites, 666, 809, 836 Stalagmites, 666, 836 Standard Light Antarctic Precipitation (SLAP), 168 Standard Mean Ocean Water (SMOW), 168 Staphylococcus, 69 Starch, degradation of, 356 Static SIMS, 883 Steranes, 152, 170, 175, 885 Precambrian, 96 Stercomata, 394 Stereocaulon vulcani, 424 Sterilization, by impact events, 70 Sterocaulon vesuvianum, 424 Sterols, 152, 170 Stichococcus, 12 Stigmastane, 152 Stigonema, 308 Stigonematales, 310 Stilpnomelane, 92 Stokes scattering, 755 Stomatocysts, 14 Storage product, in diatoms, 327 Stramenopiles, 326, 747 Stratification, 203 Stratified systems, 576 Strecker-cyanohydrin synthesis, 571 Streptococcus pneumoniae, 533 Streptomyces, 86, 151, 359 S. griseus, 151 S. pilosus, 793 Streptomycetes, 597 S. fradiae, 433 Streptophyta, 12 Streptophyta/Charophyta, 12 Stromatactis, 196, 254, 671, 847, 899 Stromatolites, 53, 97, 309, 329, 350, 617, 635, 638, 700, 809, 830 earliest, 196 endoliths in, 350 Strombus, 245 Strongylocentrotus purpuratus, 54, 55 Strontianite, 191 Strontium (Sr), 425 in carbonates, 264 isotopes (87Sr/86Sr), 516 fluid sources, 517 seawater, 517 variation in Earth history, 518 in seawater, 331 paleothermometry (Sr/Ca), 335 Subaerial exposure, of carbonates, 241 Suberan, 235 Suberites domuncula, 799, 843 Subfossil, 27 Sublacustrine hot spring, 447 springs, 475 Submarine caves, 599

SUBJECT INDEX Suboxic zone, 779 Substrate-level phosphorylation, 596 Substrates rock, 406 used by acetogens, 3 Subsurface filamentous fabrics, 851 Subterranean biosphere, 871 Subtifloria, 215 Succinite, 25 Suillus S. collimitus, 425 S. granulatus, 429 Sulfate reduction, 223, 236, 853, 859 advent, 514 alkalinity, 140 diagenesis, 780 at cold seeps, 36 role in carbonate precipitation, 223, 338 role in iron sulfide formation, 488 Sulfate–chloride waters, 470 Sulfate-methane transition zone (SMTZ), 42 Sulfate-reducing bacteria, 853, 859 Sulfate waters, 469 Sulfide, 782 formation of, 486 at mid-ocean ridges, 458 mineral oxidation, 856 oxidation, 111 at cold seeps, 283 with ferric oxides, 50 in freshwater environments, 49 in geothermal fluids, 469 in marine hydrothermal environments, 462 oxidizers, 111, 877 Sulfidic karst, 526 Sulfite, 878 Sulfobacillus, 184 S. acidophilus, 368, 372 Sulfolobus, 184, 364, 448 S. acidocaldarius, 65, 857 S. metallicus, 368 Sulfophillic stage, 902 Sulfur, 230, 457 bacteria, 877 Beggiatoa, 111 morphologically conspicuous, 878 cycle, 859 in marine hydrothermal environments, 457 isotope fractionation, mass dependent, 295 isotopes, 513 sulfur oxidizing (thiotrophic) bacteria, 87, 111, 283, 460, 877 storage, Beggiatoa, 111 Sulfurihydrogenibium azorense, 612 Sulfurimonas denitrificans, 878 Sulfurization, 150 Sulfurospirillum barnesii, 370 Supercontinent Rodinia, 206 Supernova, 569 Superoxide, 9 Superplume, 208 Supersaturation, 448, 830 CaCO3, 23 dolomite, problem, 337 Supratidal, 127 Surface charge, bacterial surfaces, 276, 614 Surfaces microbial, metal binding, 190 microbial, reactivity, 614 of rocks, 406 Surirella brebissonii, 328 Swamps, 576 Sylvite, 751

Symbiodinium, 16, 123 Symbiosis, 42, 290, 309, 356, 402, 441, 463, 675, 721, 866 corals, 252 diatoms, 329 dinoflagellates/corals, 16 foraminifera, 394 thiotrophic bacteria, 879 Synalissa, 403 Syncytium, 797 Synechobactins, 565 Synechococcus, 83, 214, 216, 307, 308, 476, 812 S. elongatus, 349, 350 Syneresis cracks, 665, 666 Synergism, 126 Syngenicity, biosignatures, 197 Synteophus, 602 Syntrophic degradation, 597 Syntrophococcus, 2 Syntrophy, 597, 866, 870 of bacteria and archaea, 39 Synurophytes, 14 Syringodium, 243 T Tank reactors, 184 Tannuolina, 61 Taphocoenosis, in amber, 34 Taphonomy, 136, 785 of Ediacaran biota, 345 Tasmanites, 789 Tellurium, 426 Temperature, on rock surfaces, 351, 406 limit for life, 364 Tepee Buttes, 849 Teponema, 84 Teredinidae, 902 Termites, 582 Terpenes, in amber, 25 Terrestrial deep biosphere, 760, 871 Terrestrial plants biomarkers, 157, compound specific isotopes, 173 rise of, 300 Tertiary oil recovery, 598 Testacealobosia, 748 Tetraether lipids, 39, 158, 885 Tetrahymanol, 173 Tetrapyrrole coenzyme, 685 Tetrapyrroles, 559 Tetrathionate, 878 Textures, 146, 833 Thalassia, 243, 245 Thalassiosira pseudonana, 802 Thalassohaline, 437, 765 Thallassinoides, 258 Thallus, 401 Thauera selenatis, 784 Thaumatomonads, 747 Thermal ionization mass spectrometry (TIMS), 503 Thermoacetogenium, 2 Thermoacidophiles, 467, 476 Thermoanaerobacter, 2 T. siderophilus, 370 Thermo-and/or halocline, 203 Thermococcus, 319, 462 T. gammatolerans, 365, 752 Thermocrinis, 82 Thermodesulfobacterium, 87 Thermodesulfovibrio, 854 Thermodynamic calculations, 460 Thermomonas, 369

SUBJECT INDEX Thermonatrite, 824 Thermophiles, 364, 460, 467, 712, 854 Thermophoresis, 708 Thermotoga, 83, 364, 448 T. maritima, 448 Thermotogales, 461, 462 Thermus, 69, 84, 319 T. thermophilus, 534 Thioalkalivibrio, 602 Thiobacilli, 878 Thiobacillus, 6, 114, 186, 476, 859, 878 T. ferrooxidans, 83, 856 T. thiooxidans, 83 Thiocapsa, 602, 687 Thiodictyon, 96, 369 Thioester World, 876 Thiomargarita, 87, 272, 296, 878 T. namibiensis, 877 Thiomicrospira, 463, 878 Thiomonas, 87 Thiophenes, meteorites, 572 Thioploca, 87, 283, 602, 859, 877, 878 Thiosulfate, (S2O23–), 462, 791, 878 Thiothrix, 283, 604, 838, 878 Thiotrophic bacteria, 877 Thiovulum, 878, 879 Thorium, 751 Thrombolites, 617, 624, 635, 830, 880 Thylakoid, 307 in cyanobacteria, 307 membrane, 736 Thyrea, 403 Tianzhushanellida, 59 Tidal flats, 880 carbonate deposits, 243 Time-of-flight secondary ion mass spectrometry (ToF-SIMS), 883 Tindallia, 2 Titan, Saturn’s moon, 77 Tolypothrix, 308 Tommotian Fauna, 63 Tommotiida, 59 Torellella, 62 Total alkalinity (TA), 20, 829 Trace fossils, 481 Neoproterozoic, 886 Traces of organisms, 189 Trametes, 420 Transition metals, Hadean ocean, 703 Transparent exopolymer particles (TEP), 361 Transvaal supergroup, 665 Trapping and binding, 329, 617, 621, 548 Travertines, 467, 470, 628, 809, 889 Trebouxia, 401, 403 T. irregularis, 406 Trebouxiophyceae, 12 Treibs, A., 148 Trentepohlia, 401 Treponema, 2 Triassic, 316 Triassic–Jurassic, extinction event, 302 Tribrachidium heraldicum, 344 Tribrachiomorpha, 345 Trichoderma, 359 Trichodesmium, 329 Trichophycus, 298 Trichosporon cerebriae, 362 Trilobita, 59 Tripanites, 125 Trona, 824 Trypanosoma gambiense, 746 Tschernozem, 835 Tsunami, 71

Tube worm, 278, 463 Riftia, 869 Tubomorphophyton, 215 Tufa, 212, 309, 889 thrombolite, 644 Tufa-forming biofilms, 309 Tumbiana Formation, 95 Tundra, 728 Tungsten, 562, 704 Typhimurium, 534 U Ultramicrobacteria, 678 Ulvophyceae, 12 Upper Ordovician, extinction event, 301 Upwelling, 203, 733, 878 Uracil, 534 Uramphite, 425 Uraninite, 752 Uranium, 751 mineralization, 693 Urease dead zones, cause for mass extinctions, 303 Urey reaction, 829 Urkingdoms, 64 Urmetazoa, 840 Uronic acids, 655 Urthiere, 747 Usnea, 404, 405 U-Th age, 415 UV photooxidation model, 96 V Vahlkampfiidae, 748 Vanadium (V), 562 van der Waals forces, 774 Vaterite, 261 Veillonella alcalescens, 84 Veins, 662, 663, 664, 665, 666 Vendobionta, 296, 345, 543 Vent. See Hydrothermal. Ventogyrus chistyakovi, 344 Vernadsky, 148 Verrucaria, 404 V. calciseda, 350 V. muralis, 352 V. rubrocincta, 350 V. rupestris, 352 Verrucomicrobia, 85, 576, 577 Vertebrate, biomineralization, 54 carcasses, 285 Vesicles, carbonate precipitation on, 224 Vesicomyid, 287 Viable count methods, 577 Vibrio, 538, 604 Vibrioferrin, 567 Vienna Pee Dee Belemnite (VPDB), 168 Vienna SMOW, 168 Viking mission, 74 Virgibacillus marismortui, 337 Viridiplantae, 10 Viruses, 767 Vital effect, 227 Vivianite [Fe3(PO4)3], 191, 780 Volborthella, 60 Volcanic, activity, 732 ash, 145 glass, 103 hot spot, 456 rocks, basalt, 103 rock weathering, 143 soils, 144

927 Volcanogenic massive sulfide (VMS) deposits, 454 Vostok, 726 W Wadden Sea, 880 Water, 75, 405 content, of rocks, 352 pore water, 742 retention, 834 -rock reactions, 457 Watsonella, 62 Waulsortian, 893 banks, 893 mudbanks, 672, 893 mud mounds, 254, 893 Weathering, 112, 143, 352, 353, 407, 732, 733, 867 of clay and silicates by fungi, 427 by endoliths, 352 of impact-shocked rocks, 72 by lichens, 407 of marine sulfides, 462 textures, 146 volcanic rocks, 143 Weddellite, 425 Wengan Phosphorites, 296 Wetlands, 581 Whale falls, 878, 901 Whewellite, 425 White rot fungi, 420, 597 White rust, 707 White smokers, 318 Whitings, 212, 219, 225, 246 Wolinella succinogenes, 83 Wood fall, 901 Wood–Ljungdahl pathway, 1 Wrinkle structures, 552 Wurtzite, 905 X Xanthoparmelia, 404 Xanthophyceae, 14 Xanthophylls, 10 Xanthoria ectaneoides, 425 Xenobiotics, 598 Xenophyophores, 394 Xeromyces bisporus, 365 Xerophiles, 365 Xylan, degradation of, 356 Xylophagainae, 902 Y Yeasts, 416 Yellowstone National Park, 350, 447, 812 Yelovichnus, 887, 888 Yorgia waggoneri, 344 Yunnanzoon, 298 Z Zebra limestones, 847 Zechstein, salts, 313 Zinc (Zn), 425, 448, 905 in marine hydrothermal environments, 457 proteins, 561 Zoochlorellae, 12 Zoophycos, 258 Zooplankton, biomarkers (compound-specific isotopes) of, 170 Zooxanthellae, 123, 252 Zygnematales, 11 Zymomonas mobilis, 84