J. Eukaryot. Microbiol., 53(4), 2006 pp. 1–13 r 2006 The Author(s) Journal compilation r 2006 by the International Society of Protistologists DOI: 10.1111/j.1550-7408.2006.00108.x
Vairimorpha disparis n. comb. (Microsporidia: Burenellidae): A Redescription and Taxonomic Revision of Thelohania disparis Timofejeva 1956, a Microsporidian Parasite of the Gypsy Moth Lymantria dispar (L.) (Lepidoptera: Lymantriidae) JIRI VAVRA,a,b MIROSLAV HYLIS,a CHARLES R. VOSSBRINCK,c DANIELA K. PILARSKA,d ANDREAS LINDE,e JAROSLAV WEISER,f MICHAEL L. MCMANUS,g GERNOT HOCHh and LEELLEN F. SOLTERi Department of Parasitology and Laboratory of Electron Microscopy, Faculty of Science, Charles University, Vinicna 7, Prague 2, Czech Republic, and b The Institute of Parasitology, Czech Academy of Sciences, Branisovka 31, Ceske Budejovice, Czech Republic, and c Connecticut Agricultural Experiment Station, 123 Huntington St., New Haven, Connecticut 06504, USA, and d Bulgarian Academy of Sciences, Institute of Zoology, 1 Tsar Osvoboditel Blvd., 1000 Sofia, Bulgaria, and e Fachhochschule Eberswalde, Alfred Moeller Str. 1, Eberswalde, D-16225, Germany, and f The Institute of Entomology, Czech Academy of Sciences, Branisovka 31, Ceske Budejovice, Czech Republic, and g USDA Forest Service, 51 Mill Pond Road, Hamden, Connecticut 06514, USA, and h Institute of Forest Entomology, Forest Pathology and Forest Protection, BOKU—University of Natural Resources and Applied Life Sciences, Hasenauerstrasse 38, A-1190 Vienna, Austria, and i Illinois Natural History Survey, 1816 S. Oak St., Champaign, Illinois 61820, USA a
ABSTRACT. Investigation of pathogens of populations of the gypsy moth, Lymantria dispar (L.) in Central and Eastern Europe revealed the existence of a microsporidium (Fungi: Microsporidia) of the genus Vairimorpha. The parasite produced three spore morphotypes. Internally infective spores are formed in the gut and adjacent muscle and connective tissue; single diplokaryotic spores and monokaryotic spores grouped by eight in sporophorous vesicles develop in the fat body tissues. The small subunit rDNA gene sequences of various isolates of the Vairimorpha microsporidia, obtained from L. dispar in various habitats in the investigated region, revealed their mutual identity. In phylogenetic analyses, the organism clustered with other L. dispar microsporidia that form only diplokaryotic spores in the sporogony cycle. The octospores of certain microsporidia infecting Lepidoptera that were previously described as Thelohania spp., have recently been shown to be one of the several spore morphotypes produced by species in the genus Vairimorpha. Because the description and drawings of a parasite described as Thelohania disparis by Timofejeva fit the characteristics of Vairimorpha, and all octosporeproducing microsporidia collected from L. dispar since 1985 are genetically identical Vairimorpha species, it is believed that the parasite characterized here is identical to T. disparis Timofejeva 1956, and is herein redescribed, characterized, and transferred to the genus Vairimorpha as the new combination Vairimorpha disparis n. comb. Key Words. Lymantria dispar, microsporidia, Nosema lymantriae, Nosema portugal, SSU rDNA sequence, Thelohania disparis, Thelohania similis, Vairimorpha lymantriae, Vairimorpha sp.
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All of the microsporidian genera recorded before 1985 were recovered in the later collections, however, P. schubergi was transferred to Endoreticulatus (Cali and El Garhy 1991), and we believe that species isolated from L. dispar and described in the genus Thelohania are actually Vairimorpha. In addition to previously described species, one collected species, Nosema portugal Maddox et al. (1999), was described thoroughly using transmission electron microscopy and molecular biology data (Table 1). In this paper, we characterize a Vairimorpha sp. from L. dispar found in Rupite, Bulgaria in 1995. This isolate was chosen among others collected because the germplasm was isolated from a single L. dispar larva collected in a specifically documented site. The pathogen was monitored in the host population for 15 yr and the biology of this isolate was elucidated in several previous studies (Pilarska 1987; Pilarska et al. 1998; Solter and Maddox 1998a).
ICROSPORIDIA isolated between 1927 and 1985 from populations of the gypsy moth Lymantria dispar (L.) in Central and Eastern Europe were placed in the genera Pleistophora (formerly Plistophora), Thelohania, and Nosema (Timofejeva 1956; Weiser 1957, 1961; Zwo¨lfer 1927a, b). While three of the described isolates, Nosema lymantriae, Nosema serbica, and Pleistophora schubergi figure as valid species, the taxonomic status of isolates reported from mixed infections, those assigned to the genus Thelohania, and those that were reported cross-infecting populations of L. dispar and Euproctis chrysorrhoea (L.), the browntail moth (Weiser 1957; Table 1) remain uncertain. A group of European and U.S. scientists collected more than 30 microsporidian isolates from populations of L. dispar in Austria, Bulgaria, Czech Republic, Germany, Hungary, Poland, Portugal, Romania, and Slovak Republic from 1985 to 2002. These microsporidia are stored in the liquid nitrogen collection of the Illinois Natural History Survey under United States Department of Agriculture/Animal and Plant Health Inspection Service/Plant Protection Quarantine (USDA/APHIS/PPQ) permit and are studied for their potential use as classical biological control agents against L. dispar in the United States where gypsy moth microsporidia do not occur (Campbell and Podgwaite 1971; Podgwaite 1981), and for augmentative release in Europe.
MATERIALS AND METHODS Origin of the microsporidian isolates. An isolate of a Vairimorpha-type microsporidium was obtained from L. dispar larvae feeding on Salix alba L. in Rupite, Bulgaria (23116 0 1100 East, 41128 0 5600 North) in 1995. Tissues from the infected larva were homogenized and filtered. The spores were cleaned in several tap water washes using centrifugation, stored in a 1-ml suspension of distilled water containing 5 mg streptomycin and 400 U fungizone per ml suspension, and were transported to the Center for Ecological Entomology, Illinois Natural History Survey, Champaign, Illinois, where they were stored in liquid nitrogen (INHS Accession number 1995-D). This microsporidium was recovered from
Corresponding Author: L. Solter, Center for Ecological Entomology, Illinois Natural History Survey, 1816 S. Oak St., Champaign, IL 61820—Telephone number: 217-244-5047; FAX number: 217-3334949; e-mail:
[email protected]
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Table 1. Described species of microsporidia from field populations of Lymantria dispar and isolates from Bulgaria that are relevant to these descriptions. Origin of isolate Germany
Identification Pleistophora schubergi schubergia,b Thelohania disparisa Nosema lymantriaea Nosema muscularisa
Tissues infected Midgut
Vajnory, Bratislava-Dvornik, Slovakia (1955) Serbia, Yugoslavia, Varna, Bulgaria Bulgaria (recovered by Pilarska, 1984) Rupite, Bulgaria
Thelohania similisa
Fat body Silk gland, fat body Circular & longitudinal muscle of midgut Fat body
Nosema serbicaa
Various
N. lymantriae & T. similis Vairimorpha sp.
Fat body
Levishte, Bulgaria Asenovgrad, Bulgaria Lisbon, Portugal
Nosema sp. Endoreticulatus sp. Nosema portugala
Ukraine Sv. Benadik, Slovakia (1955) Sv. Benadik, Slovakia (1955)
Fat body silk gland & fat body midgut Silk gland & fat body
Remarks/References Plistophora (Zwo¨lfer, 1927a, b) Pleistophora (Kaya 1973) Timofejeva (1956) Weiser (1957) Weiser (1957) Now considered primary spores of Nosema & Vairimorpha spp. (Weiser and Linde 1998). Weiser (1957); from Euproctis chrysorrhoea Weiser (1961), Sidor (1979) Pilarska and Vavra (1991) David and Weiser (1989) Probably Vairimorpha based on TEM & sequence data Pilarska et al. (1998) Reported in Ph.D. thesis as Nosema1Thelohania Pilarska (1987) Pilarska et al. (1998) Probably E. schubergi; Wang et al. (2005) Maddox et al. (1999)
a
Described species. Endoreticulatus schubergi (El Garhy and Cali 1991) described from Choristoneura fumiferana. TEM, transmission electron microscopy. b
L. dispar populations at the same site over a period of years (Pilarska et al. 1998; Solter, Pilarska, and Vossbrinck 2000). Inoculation of L. dispar larvae. Third-instar Lymantria dispar larvae (New Jersey Standard strain, USDA/APHIS Otis Method Development Center, Cape Cod, MA) were fed 3 105 spores (both diplokaryotic spores and octospores) on small diet cubes. Larvae that completely consumed the diet blocks were reared on high wheat germ diet (Bell et al. 1981) in growth chambers at 24 1 1C, 16:8 h L:D and were dissected at intervals ranging from 12 h to 20 days post-inoculation (pi). Microscopic studies. Freshly dissected tissues were examined under phase contrast microscopy at 400 and 1,000. Tissue smears were stained either with Giemsa (Sigma Diagnostic Accustain) or with Giemsa after acid hydrolysis in 1 N HCl (Vavra and Maddox 1976). Agar-immobilized spores (Vavra 1964) (n 5 100 per spore type) were measured using the LeicaTM Image Analysis System (Pilarska, Linde, and Pilarska 2000). For transmission electron microscopy (TEM), infected tissues were either fixed for 24–48 h in 2.5% (v/v) glutaraldehyde in 0.1 M cacodylate buffer, pH 7.0, and post-fixed in 2% (w/v) aqueous OsO4, or were fixed for 1 h in 0.1 M phosphate buffer containing 1% (v/v) glutaraldehyde and 1% (w/v) OsO4 (Hirsch and Fedorko 1968). Fixed tissues were washed in the respective buffers used for fixation, dehydrated through ascending ethanol and acetone series, and embedded either in Epon-Araldite or in Poly/ Bed 812/Araldite 502 (Polysciences Inc., Eppelheim, Germany). Thin sections were cut on a Reichert OMU2 and Ultracut E ultramicrotomes, stained with uranyl acetate and lead citrate, and examined with Zeiss EM9S and JEOL 1010 electron microscopes. To determine development times and proliferation of parasites in the tissues, infected larvae (dosage 5 103 spores in thirdinstar larvae; 24 1 1C; 16:8 h L:D) were dissected every 24 h, 24–144 h, then 240 h pi. Fresh midgut and fat body tissues were individually spread on glass microscope slides and pressed under glass coverslips to a one-cell thickness. Spores and other developmental forms were counted per 525 or 1,000 phase contrast microscopic field, 81 total sample counts of anterior, center, and posterior portions of each tissue from each of two or three larvae per time period pi. DNA isolation, amplification, sequencing and sequencing analysis. Fat body tissues collected from L. dispar larvae infected
for 21 days were filtered through nylon mesh cloth, centrifuged to pellet the spores, and purified on a 1:1 sterile water:Ludoxs HS40 colloidal silica gradient (Sigma-Aldrich, St. Louis, MO), a total of 35 ml in 50-ml plastic centrifuge tubes. The spores were cleaned by centrifugation in sterile water and delivered on ice packs to the Connecticut Agricultural Experiment Station for sequencing preparation. A 10-ml suspension of spores was pelleted by centrifugation, resuspended in 150 ml TAE buffer (0.04 M Trisacetate, 0.001 M EDTA) in 0.5-ml micro-centrifuge tubes, shaken with 150 mg of 0.5 mm glass beads in a Mini-Beadbeater (Biospec Products, Bartlesville, OK), and incubated at 95 1C for 3 min (Vossbrinck et al. 2004). One to three microliters of the TAE-ruptured spore suspension were removed and a standard PCR reaction was conducted (94 1C for 3 min, followed by 35 cycles of 94 1C for 45 s, 45 1C for 30 s, and 72 1C for 90 s) using primers 18f and 1492r. The PCR product, 1,200 nucleotides in length, was purified using a Qiaquick PCR purification kit (Qiagen Company, Valencia, CA). Sequencing was performed at Keck Biotechnology Resource Laboratory at Yale University using the following microsporidian primers: 18f, 5 0 –CACCAGGTTGATTCTGCC-3 0 ; SS350f, 5 0 -CCAAGGA(T/ C)GGCAGCAGGCGCGAAA-3 0 ; 350r, 5 0 -CCGCGG(T/G)GCT GGCAC-3 0 ; 1047r, 5 0 -AACGGCCATGCACCAC-3 0 ; 1061f, 5 0 GGTGGTGCATGGCCG-3 0 ; and 1492r, 5 0 -GGTTACCTTGTTA CGACTT-3 0 . The ClustalX program (Thompson et al. 1997) was used to align sequences, which were compared to L. dispar microsporidia sequences currently deposited in GenBank and analyzed using PAUP version 4.0b10 (Swofford 1998). Bootstrap analyses using both maximum parsimony and neighbor joining search methods were performed using PAUP Version 4.0b. One-thousand replicates were run for each method. All characters were given equal weight. Amblyospora californica was assumed to be an outgroup to the Vairimorpha/Nosema clade (Vossbrinck and DebrunnerVossbrinck 2005). RESULTS Light microscopy. The microsporidium has a complex life cycle, characterized by three sporulation modes that occur in different tissues and result in spores of different structure. The
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sporulation modes are the primary cycle (S1), the secondary single-spore cycle (S2ss), and the secondary octosporous cycle (S2oct). Each cycle is comprised of a merogonial multiplicative phase (structurally indistinguishable in the respective cycles) followed by a sporogony typical for the individual cycles. The gut epithelia and, more commonly, gut longitudinal and transverse muscle cells are the sites for the primary cycle and the fat body is the principal target tissue for the secondary cycles. Other tissues, including gonads, can occasionally be infected (see Discussion). Light microscopy, primary sporulation (S1) cycle. The S1 cycle produces thin-walled, binucleate, single diplokaryotic spores (the primary spores). Merogonial stages of microsporidian development in midgut cells of the L. dispar host are indistinguishable from those of the closely related N. portugal (Maddox et al. 1999) and other Nosema-type isolates recovered in our surveys. The infection begins when ingested secondary spores inject sporoplasms, small (1.5 mm) binucleate cells (Fig. 1), into gut epithelial cells, gut muscle cells, and connective tissue. Some sporoplasms are also injected into hemocytes adhering to the gut and to tracheal matrix cells. While growing, the sporoplasms (Fig. 2) transform into meronts, larger cells (2.5–4.0 mm) with two nuclei in the diplokaryon configuration (Fig. 3). The meronts grow, their nuclei divide, and larger cells of approximately 10 mm with two diplokarya are formed (Fig. 4–6). These cells divide into binucleate forms and the merogonial cycle is repeated. Occasionally, cells with a large, seemingly single nucleus are found in Giemsa smears (Fig. 7). Presumably after several divisions, the binucleate meronts mature into binucleate sporonts in which the nuclei divide forming tetranucleate (two diplokarya) sporonts (7–10 mm) (Fig. 8–11). Each tetranucleate sporont divides once (Fig. 12), giving rise to two binucleate sporoblasts (bisporous sporogony) in a chain of approximately 3 11 mm (Fig. 13). The sporoblasts separate (Fig. 14) and mature individually into spores that lie free in the host cell cytoplasm. Mature primary spores are elongate-ellipsoidal, measuring when fresh 5.4 0.48 2.5 0.37 mm, have thin walls (i.e. are less refractive under phase contrast), and have a large posterior vacuole (Fig. 15). The spores spontaneously extrude their polar filaments while inside the host tissue. It is not known whether the sporoplasms repeat the primary sporulation cycle or begin the secondary cycle in its two modes. The S1 cycle appears to repeat, at least to a modest extent, in other tissues, including the fat body. Secondary single spore (S2ss) cycle. The secondary cycle takes place almost exclusively in the fat body cell cytoplasm. The S2ss cycle produces thick-walled, binucleate, secondary spores of the Nosema type. The early stages cannot be distinguished structurally using light microscopy from those of the primary cycle, except that the S2ss meronts and S2ss sporonts are found in fat body cells. The spores produced are elongate ellipsoidal, measure when fresh 5.1 0.34 2.6 0.28 mm, are more refractive than the primary spores, do not show clearly the pos-
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terior vacuole (Fig. 16), and display a prominent diplokaryon (Fig. 22) after staining. Secondary octosporous cycle (S2oct). Thick-walled, uninucleate secondary spores of the Thelohania type (octospores) are formed in this cycle. Merogonial stages are structurally indistinguishable from those of other cycles but, as in the case of the S2ss cycle, they occur in fat body cells. The first recognizable stages of the S2oct cycle are binucleate sporonts with one diplokaryon (Fig. 17) that later separates into two independent nuclei (Fig. 18). The sporonts, approximately 10 mm in diam., are characteristically rounded in outline, suggesting that a sporophorous vesicle (SV) is being formed around them (Fig. 17–21). The two independent nuclei divide once, forming a cell containing four single nuclei (Fig. 20). In Giemsa smears, the nuclei of the octosporous sporulation stages typically have a fuzzy appearance (Fig. 19). Karyokinesis and cytokinesis then give rise to eight sporoblasts that mature into spores enclosed within a semi-persistent SV 7.3– 10.5 m in size (Fig. 21–22, 24–25). The spores are broadly oval, 4.6 0.28 2.8 0.26 mm in size when fresh, are highly refractive under phase contrast, and seemingly have no posterior vacuole (Fig. 25). Stained spores show one dot-like nucleus (Fig. 22). For an unknown reason, the karyokinesis distributing nuclei into individual spores fails in some SV. Such SVs contain fewer spores and some remaining spores may have two nuclei instead of one. The total number of nuclei in the SV, however, is always eight (Fig. 23). Ultrastructure. Although the microsporidium has a fairly complex life cycle consisting of three different sporulation modes, the only differences in the fine structure of individual sporulation cycles are those of late sporonts, sporoblasts, and spores. Individual merogonial stages cannot be attributed to individual sporulation modes with certainty using fine structure as the sole criterion. S1 and S2ss cycles. Development begins when a sporoplasm is injected by an ingested secondary spore or by a primary spore into a suitable host cell. The injected sporoplasm is a small, simple cell, enveloped by a single plasma membrane, with two nuclei in imperfect diplokaryon configuration. Ribosomes are scattered throughout the cytoplasm. Endoplasmic reticulum (ER) or Golgi vesicles are missing (Fig. 26). Host cell ribosomes assemble on the outer face of the sporoplasm plasmalemma (Fig. 27). The sporoplasm grows into a meront characterized by the presence of a few endoplasmic reticulum membranes and an increased number of cytoplasmic ribosomes (Fig. 28). As shown using light microscopy, meronts grow into tetranucleate stages and divide into binucleate forms again. This process is evidently very rapid and this stage was not documented in TEM sections. The transition of meronts into sporonts, the initial stage of the sporogony cycle, follows a pattern typical for microsporidia. The meront–sporont transition stage is diplokaryotic and contains numerous ER cisternae. The plasmalemma is thickened by a thin layer of dense material that first occurs in patches and later circumscribes the cell. Because the ER cisternae are more abundant at the onset of
Fig. 1–25. Vairimorpha disparis developmental stages and spore morphotypes as seen under light microscopy. Scale bar 5 5 mm in all figures. 1–14, 18, 19, 21 smears stained by Giemsa; 17, 20, 23 smears stained by Giemsa after 1 N HCl hydrolysis; 15, 16, 24, 25: fresh spores. 1. Sporoplasm of the primary sporulation (S1) mode (arrow). 2. Mature sporoplasm 3. Early meront. 4. Group of meronts of various sizes with diplokaryotic nuclei. 5. Late meront with dividing diplokaryon. 6. Meront with four nuclei. This is the maximum number of nuclei formed in a cell. 7. Cell with a large, apparently single nucleus. 8. Late meront/early sporont with two closely adjacent diplokarya. 9–11. Late meronts/early sporonts with two diplokarya prepare for cell division. 12. Dividing sporont. 13. Sporont division yields two sporoblasts in a chain. 14. Individual sporoblast will mature into a diplokaryotic spore. 15. Spores of the S1 cycle formed in the cells of the tracheal matrix. Note the large posterior vacuoles (Nomarski interference contrast). 16. Spores of the secondary single-spore sporulation (S2ss) cycle (Phase contrast). 17–21. Sporonts of the secondary octosporous sporulation (S2oct) cycle. 17. Two early sporonts with nuclei still in the diplokaryon configuration. 18. Late sporont in which the diplokaryon nuclei have separated. 19. Late sporont with separated nuclei have begun meiosis. Note the characteristic fuzzy appearance. 20. Meiosis results in a plasmodium with four nuclei. 21. Mitosis following meiosis yielded an eight-nucleate plasmodium. 22. Sporophorous vesicle (SV) with eight mature, uninucleate octospores, with diplokaryotic S2 single spores (arrow) in its vicinity. 23. SV, product of a sporont in which nuclear separation during spore formation partly failed. There are six spores in the SV instead of eight, two of them aberrant, having two nuclei. 24, 25. SVs with immature (Fig. 24) and mature (Fig. 25) octospores.
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sporulation and the ribosomes assemble on ER, the cytoplasm of early sporonts appears more transparent than those of meronts (Fig. 29–30). Sporoblasts of both the S1 and S2ss cycles are very dense, heavily wrinkled cells, lying freely within the host cell
cytoplasm. Only the diplokaryon can be recognized (Fig. 31). S1 spores can easily be distinguished from spores of both secondary cycles. They have a thin exospore and endospore, and four to five (rarely six) polar filament coils arranged in a single row
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(Fig. 32–34). The area of posterior vacuole is characteristically collapsed in TEM preparations of mature S1 spores (Fig. 34). The S2ss spores usually have 11–13 polar filament coils arranged in one row (Fig. 35–36). Mature S2ss spores have a thick (120– 150 nm) endospore and slightly wrinkled exospore of approximately 30 nm. Because of the dense spore contents, no other organelles were observed in mature S2ss spores. The octosporous sequence. At the beginning of the S2oct sporulation, sporonts are similar in shape to those of S1 and S2ss (compare Fig. 30 and 37). During development, however, these sporonts form a sporophorus vesicle on their surface. The SV membrane appears on the sporont plasmalemma in small, blisterlike patches in which dense material is being deposited (Fig. 37, 38). Sporonts at this stage of development are difficult to fix and the cytoplasm and cell membrane are not well defined (Fig. 37). As development proceeds, the sporonts become rounded in shape and the SV appears as a thin membranous layer loosely ensheathing the sporonts (Fig. 39). In the episporontal space thus formed, patches of very dense granulo-fibrilar material are secreted by the sporonts, forming a tortuous mesh (Fig. 38, 39, 41–48). This material evidently originates in the sporont cytoplasm as small, circular vesicles containing dark material and is exported by Golgi vesicles (Fig. 43). When the development of the SV is complete, the sporont contains an elaborate system of ER cisternae arranged in circular fashion around the two nuclei, which adhere less tightly to each other (Fig. 39) than in a diplokaryon (Fig. 37). Dark granular or rod-like patches resembling synaptonemal complexes occur in the nuclei at this stage (Fig. 39, 40). This stage is very fixation sensitive, probably due to permeability properties of the SV, which tends to swell. Thus, the sporont plasmalemma is usually broken in some places (Fig. 39). During sporogony, the two sporont nuclei become fully separated (Fig. 41). The nuclei divide and a four-nucleate plasmodial stage is formed (Fig. 44). The karyokinesis and cytokinesis that follow give rise to eight uninucleate daughter cells inside the SV (Fig. 45, 47). This developmental pattern and the occurrence of synaptonemal-like structures in the nuclei seem to indicate that the diplokaryotic nuclei of the sporont undergo a two-step meiosis after separation. During the advancing sporogony, the layer of secretory material in the episporontal space becomes more prominent, filling the entire volume of the SV (Fig. 45, 47). The cell inside the SV links with the dense secretory material in the SV lumen by numerous plasmalemma protrusions (Fig. 42). The sporont daughter cells gradually develop into sporoblasts by deposition of a thick layer of dense material (the future exospore) on the cell membrane and by intracytoplasmic differentiation during which the cytoplasm and its structures become polarized. A large vacuole, sometimes appearing in TEM sections as two independent vacuoles because of its tortuous shape, is formed at one pole of the cell. A Golgi network of vesicles and channels forms the primordium of the polar filament at the opposite side of the cell (Fig. 48). The vac-
uole mentioned above is probably the primordium of the polaroplast. Dense membrane whorls observed in some sporoblasts seem to originate from the vacuole by membrane infolding (Fig. 46). While the sporoblast matures, the cytoplasm becomes more dense, and the cell outline more tortuous. At the same time, the amount of the dense secretory material inside the SV diminishes (Fig. 49) and nearly disappears in the mature SV. Each sporoblast develops into one spore. Some internal details can be seen only in immature octospores, as mature octospores are so dense and poorly preserved that only polar filament coils can be seen in their interior. The young octospore has a single nucleus, many polaroplast lamellae, a few ER cisternae, and a large Golgi area situated posterior to the nucleus. The approximately 30 polar filament coils are arranged partly as a single layer and partly as double or triple layer. A very thick, electron-dense layer, the future exospore, covers the cell (Fig. 50). Mature octospores have a very thick exospore (around 100 nm), very thick endospore (120– 150 nm), and 28–30 polar filament coils arranged in an irregular double layer (Fig. 51). As described previously, the mechanism distributing the postmeiotic nuclei into individual spores fails in some SV and, rarely, sporoblasts and young spores with two nuclei can be found inside SV (Fig. 52, 53). Sequence data. The dataset for the small subunit rDNA has a total of 2,060 characters; 1,319 were constant and 195 were parsimony informative. Both parsimony and neighbor-joining methods gave identical topologies (Fig. 54) with 100% bootstrap values at all but two branch points. The ssrDNA sequences obtained for the Rupite, Bulgaria isolate 1995-D characterized above were compared to the following Vairimorpha spp. isolated from L. dispar: isolate 2004-A collected from the same site 9 years later; isolate 1995-C collected in Bulgaria at approximately 1 km distance from the collection site of 1995-D; isolate 1995-B from Slovakia; and isolate 1985-B from Czech Republic. The sequences of all Vairimorpha isolates were identical and are closely related to N. portugal and N. lymantriae, both representatives of the group of gypsy moth microsporidia forming solely single, diplokaryotic spores in the secondary sporogonic cycle (Fig. 54). DISCUSSION Most of the life cycle of the microsporidium described here is indistinguishable from that of N. portugal (Maddox et al. 1999) and other Nosema-like (i.e. single-spore forming) isolates recovered in our surveys from the L. dispar populations (McManus and Solter 2003) except that octospores are produced in the S2oct cycle and secondary spores are typically absent or do not mature in the silk glands. The occurrence of two spore types in the S2 cycle, diplokaryotic single spores and monokaryotic octospores, places this species in the genus Vairimorpha as defined by Pilley (1976). Thus far, no valid taxonomic description exists for a Vairimorpha sp. from the gypsy moth.
Fig. 26–36. Fine structure of Vairimorpha disparis primary (S1) and secondary single spore sporulation (S2ss) cycles. 26. Sporoplasm soon after being injected into the host cell. The cytoplasm contains some free ribosomes, the endoplasmic reticulum (ER) and Golgi are not yet formed, but the nuclei (N) are already in an incomplete diplokaryon configuration and contain nucleoli (). Scale bar 5 500 nm. 27. Detail of Fig. 26. Host cell ribosomes assemble around the sporoplasm. Scale bar 5 500 nm. 28. Early meront characterized by numerous cytoplasmic ribosomes, poorly developed ER and the absence of Golgi vesicles. Scale bar 5 500 nm. 29. Group of meronts and young sporonts. Sporonts (S) are distinguished from meronts (M) by thicker cell membrane, paler cytoplasm and more abundant ER. Scale bar 5 1 mm. 30. Sporont of the S1 or of the S2ss cycle. The electron-dense material, characterizing the meront to sporont conversion, covers most, but not yet entire, cell membrane. Scale bar 5 1 mm. 31. Sporoblast of the S1 or of the S2ss cycle. The cell is electron-dense, is characteristically wrinkled, and only the two nuclei (N) are visible. Scale bar 5 500 nm. 32. Immature spores of the S1 cycle. Note four to five coils of the polar filament and two nuclei (N), the relatively thin exospore adhering loosely to the cell and the thin endospore. Scale bar 5 500 nm. 33. Polar filament is arranged in 5 (range 4–6) coils in the primary spore. Scale bar 5 200 nm. 34. Mature primary spore with two nuclei (N), 4/5 coils of the polar filament, a thin exo- and endo-spore. The area of the posterior vacuole is characteristically collapsed in TEM processed primary spores. Scale bar 5 1 mm. 35. Young single-spores of the S2ss cycle with 11–12 polar filament coils, Golgi vesicles (G) and a nascent polaroplast (arrow). Scale bar 5 500 nm. 36. Mature, single-spores of the S2ss cycle showing nuclei (N) and 12–13 polar filament coils. Compare the thickness of the exospore (EX) and endospore (EN) and the number of polar filament coils with those of primary spore in Fig. 34. Scale bar 5 500 nm.
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A microsporidium from L. dispar forming eight spores within a SV (octospores) was first reported by Timofejeva (1956), who described an isolate, probably collected in the Ukraine during an L. dispar outbreak (Weiser 1998), as Thelohania disparis, n. sp. Timofejeva (1956) did not mention the occurrence of single spores, but one of her drawings indicates that she observed both octospores and single spores (Fig. 55). Hazard and Oldacre (1975) expressed doubt that T. disparis was a member of the Thelohaniidae, although they did not elaborate their reasons. In addition to Timofejeva’s report of octosporous microsporidia in L. dispar, Weiser (1957) described a second such species he named Thelohania similis. This species, however, was described from the browntail moth, Euproctis chrysorrhoea, but was reported in the same paper to have been found infecting L. dispar in Slovakia in ‘‘mixed infections’’ with what was believed to be Nosema lymantriae, a microsporidian simultaneously described from L. dispar (Weiser 1957). We believe that Weiser’s mixed infection in L. dispar was actually a Vairimorpha infection and that the spores believed to be those of N. lymantriae were the single-type secondary spores of Vairimorpha. In this respect, Vairimorpha necatrix, the type species of the genus Vairimorpha, was believed to be a mixed species infection of two microsporidia, Nosema necatrix and Thelohania diazoma (Maddox and Sprenkel 1978; Pilley 1976). We conclude that N. lymantriae (Weiser 1957), which produces single diplokaryotic spores but not octospores, is a legitimate species because Weiser’s description of N. lymantriae reported infections that produced only single spores predominantly in the silk glands. Nosema muscularis Weiser is now recognized to be the S1 cycle primary spores of both the Nosema spp. and Vairimorpha isolates of L. dispar (Weiser and Linde 1998). Pilarska (1987) reported a mixed infection, similar to those of Weiser (1957), from a Bulgarian L. dispar population and referred to the isolate as a mixed infection of N. lymantriae and T. similis (INHS isolate 1985-C). Ultrastructural drawings and the characterization of this isolate produced by David and Weiser (1989) are nearly identical to the photographs produced for this current study, but polar filament coils numbered five, a condition we found only in the primary cycle (S1) of the Nosema-type spores. The Bulgarian isolate was subsequently recognized to be a Vairimorpha species (Pilarska et al. 1998) and a 1995 isolate from the same site was studied for this description. Originally assumed to be the species represented by the reported mixed infections of N. lymantriae and T. similis, the names N. lymantriae and Vairimorpha lymantriae were used provisionally in some published reports (Linde, Genthe, and Lacker 1998; Maddox et al. 1999; Solter, Maddox, and McManus 1997; Weiser and
Novotny 1987) and in GenBank submissions (Table 2). We choose to treat the Vairimorpha species characterized in this paper as identical to the microsporidium reported by Timofejeva (1956) for the following reasons: (1) N. lymantriae is a legitimate species (Weiser 1957); (2) T. similis mixed with N. lymantriae was described from E. chrysorrhoea rather than L. dispar (Weiser 1957); (3) The relationship between the Vairimorpha microsporidia from E. chrysorrhoea and L. dispar is unknown. As the organism is clearly different from all described Vairimorpha species, we redescribe and characterize the species originally described by Timofejeva (1956) as Vairimorpha disparis (Timofejeva) n. comb. The microsporidium described here is clearly different from other species of the genus Vairimorpha infecting Lepidoptera (Table 3). The differences concern host specificity and structural and molecular characters. The following Vairimorpha species have been described from lepidopteran hosts: V. necatrix (Kramer 1965), V. plodiae (Kellen and Lindegren 1968), V. ephestiae (Mattes 1928), V. imperfecta (Canning et al. 1999), V. mesnilli (Paillot 1918), and V. heterosporum (Kellen and Lindegren 1969). Additionally, Nosema antheraeae Simchuk, Lysenko, and Tchetkarova, 1979, described from Antherea pernyi (Gue´rinMe´neville 1855) (Simchuk et al. 1979) has been referenced as a Vairimorpha (Yefimenko, Sokolova, and Issi 1990), although it has never been redescribed (Sokolova, pers. commun.). Microsporidia from Lepidoptera have spores generally similar in size and shape to each other and to the microsporidium described here. Some ultrastructural data enabling an in-depth comparison are available for the first four of the above-mentioned species. The type species of the genus, V. necatrix (Kramer 1965), although of low host specificity, is clearly a different species. The number of polar filament coils is different for spores produced in the S2ss and S2oct cycles, and secretory products in the episporontal space are lamellar rather than web-like (Moore and Brooks 1992). Differences are also found among other lepidopteran Vairimorpha species (Table 3). All sequenced Vairimorpha species are distinct from V. disparis. Molecular relationships and phylogenetic position of V. disparis. Phylogenetically, Vairimorpha isolates and the related N. portugal and N. lymantriae form a highly supported clade separate from Vairimorpha necatrix, the type species of the genus Vairimorpha (Baker et al. 1994; Vossbrinck et al. 2005). A microsporidium entered in the Genbank database as Vairimorpha lymantriae (AF141129), but forming only single diplokaryotic spores, was provisionally named Nosema sp. [Levishte isolate] (Fig. 54); it is comparable to other Nosema-like isolates from
Fig. 37–48. Fine structure of Vairimorpha disparis secondary octosporous sporulation (S2oct) mode. 37. Early sporont. The electron-dense material of the future sporophorous vesicle (SV) detaches as a thin layer from the electron-dense material covering the plasmalemma of the sporont (arrows). The nuclei are in the diplokaryon configuration (N,N) and contain nucleoli (). The cytoplasm contains a number of ribosomes and small fragments of endoplasmic reticulum. Scale bar 5 500 nm. 38. Detailed view of the early sporont (SP) border zone with the host cell (H). The plasma membrane of the microsporidian is patchily covered by electron-dense material (dark arrow). The overlaying electron-dense material (white arrow) represents the SV wall. Electron-dense secretory products are deposited, initially as small patches in the nascent episporontal space. Scale bar 5 500 nm. 39. Advanced sporont. The SV membrane (arrow) is now fully separated from the plasmalemma (arrowhead), electron-dense secretory products () start to appear in the episporontal space (ES). Endoplasmic reticulum is now fully developed, the two nuclei of the former diplokaryon separate and a synaptonemal like structure appears in one of them. Scale bar 5 500 nm. 40. Detailed view of one of the two nuclei from Fig. 41 showing the synaptonemal-like complex structure (arrow). Scale bar 5 500 nm. 41. Composite figure showing various stages (a,b,c) of diplokaryon separation into individual nuclei which occurs in the sporont at the onset of sporulation. Note the dark, filamentous material within the separating nuclei. Scale bar 5 500 nm. 42. Part of the late sporont showing a layer of numerous vesicular protrusions of the parasite plasmalemma (), mediating the contact of the parasite with the secretory material in the form of irregular electron-dense meshwork. Scale bar 5 500 nm. 43. In the advanced sporont the secretory products are exported from the Golgi area throughout the cytoplasm in the form of membrane-bound vesicles with dense central spot. Scale bar 5 500 nm. 44. Multinucleate plasmodium is formed within the SV after nuclear separation and the first meiotic nuclear division. Scale bar 5 500 nm. 45. Progressive separation of sporont daughter cells into individual cells, future sporoblasts and spores. Nascent polaroplast is at arrow. Scale bar 5 1 mm. 46. The polaroplast is formed by progressive infolding of a cytoplasmic vacuole (see in Fig. 48). Scale bar 5 250 nm. 47. SV in which the cytokinesis is completed and the cytoplasmic differentiation into extrusion apparatus organelles of future spores takes place. The whole volume of the SV is now filled with the dense threads of the secretory material. Scale bar 5 1 mm. 48. Detailed view of one cell from Fig. 47 showing the Golgi (G) forming the first polar filament coil (arrow) and a large vacuole (), probably the primordium of the polaroplast. Scale bar 5 1 mm.
VAVRA ET AL.—VAIRIMORPHA DISPARIS N. COMB. IN LYMANTRIA DISPAR
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Fig. 49–53. Fine structure of Vairimorpha disparis secondary octosporous sporulation (S2oct) cycle. 49. Sporoblast maturation involves the progressive disappearance of the electron-dense secretory material from the sporophorous vesicle (SV) lumen. Scale bar 5 500 nm. 50. Young octospore possessing about 30 polar filament coils, a single nucleus (N) and some polaroplast lamellae (arrow). The electron-dense exospore has reached its final thickness, but the endospore is absent. Scale bar 5 500 nm. 51. Part of mature octospore demonstrating the thick endospore (EN) and exospore (EX) and about 28 coils of the polar filament arranged mostly as a double row. Scale bar 5 500 nm. 52, 53. Two SVs in which nuclear separation during sporogony failed, resulting in the formation of aberrant binucleate (N,N) sporoblasts. Polar filament primordium (future anchoring disc) is at thick arrow (Fig. 53), nascent polar filament coils in Fig. 52 and 53 are at thin arrows. Scale bar 5 500 nm.
L. dispar. This species might, in fact, be a Vairimorpha that has lost the octosporous sporogony sequence. Canning et al. (1999) reported the existence of such a process for Vairimorpha imperfecta, a parasite of Plutella xylostella (L.). It thus appears that octospore formation may be an ancestral character and that the genera Nosema and Vairimorpha should, at minimum, be placed in the same family (Canning et al. 1999). Our research results and the molecular data obtained on microsporidia of L. dispar (Baker et al. 1994; Canning et al. 1999; Maddox et al. 1999; Vossbrinck, et al., unpubl. data; and this paper) suggest that isolates from L. dispar having a Vairimorpha-like cycle and those having a Nosema-like cycle, are closely related phylogenetically and could represent either different species of the same genus or perhaps even a single species based on ssrDNA or lsrDNA sequences. Baker et al.
(1994) suggested that the presence/absence of a diplokaryon or a SV might have less phylogenetic significance at higher taxonomic levels than previously thought. Major revision of the conventional classification system of microsporidia is, however, beyond the scope of the present paper. Biology and ecological interactions of V. disparis. The biology of this Vairimorpha species and its interactions with the host and with other parasitic species have been reported by a number of researchers since 1985. Pilarska (1987) reported host effects of the pathogen. Macroscopically, the infected fat body appears white and swollen. The host cell cytoplasm is filled with diplokaryotic spores and octospores. Cell membranes, however, remain intact even in the final stages of infection. Heavily infected L. dispar larvae fail to pupate, or succumb to abnormal pupation.
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J. EUKARYOT. MICROBIOL., VOL. 53, NO. 4, JULY– AUGUST 2006 Amblyospora californica Vairimorpha sp. (from Solenopsis richteri ) Vairimorpha cheracis Vairimorpha imperfecta Nosema bombycis Vairimorpha necatrix Nosema portugal Vairimorpha disparis Nosema sp. "Levishte" Endoreticulatus schubergi Endoreticulatus bombycis
Fig. 54. A phylogenetic tree of microsporidian species showing the position of Vairimorpha disparis n. comb. based on maximum parsimony (MP) analysis using the branch and bound search method. Maximum likelihood and neighbor joining (NJ) analyses gave identical topologies. The numbers at the nodes are the bootstrap value of 100 replicates for MP and 1,000 replicates for NJ.
immature diplokaryotic spores are found in the silk glands and in midgut transverse and longitudinal muscle tissues, no mature spores or octospores have been observed in these tissues. Formal ID50 and LD50 studies were not conducted, but bioassays performed at the INHS laboratory produced infection in 100% of hosts when mid-third instar L. dispar larvae were inoculated with 100 spores. At dosages between 100 and 5,000 spores, mortality usually occurred during the final stadium or during pupation. Hoch and Schopf (2001) observed 100% larval mortality when L3 larvae were inoculated with 1,000 spores per larva using the genetically identical 1985-B isolate, and at dosages above 105 spores, larvae died within a few days of inoculation. Transmission appears to be oral via infective spores disseminated in decomposing cadavers, but a small amount of horizontal transmission occurred among living infected and uninfected larvae in laboratory bioassays (Solter and Maddox 1998b). Although gonads are infected, vertical transmission has not been shown in laboratory studies and larvae infected at third instar with dosages as low as 100 spores do not eclose as adults. Further studies are needed to determine whether the pathogen is vertically transmitted. Pilarska et al. (1998) recorded the occurrence of Vairimorpha infection in L. dispar for 10 seasons of a 14-year period in the type locality. Prevalence ranged from 1% to 30% of the late instar population. The host range of the L. dispar Vairimorpha was studied in North American forest Lepidoptera (Solter et al. 1997). Laboratory infections indicate that the microsporidium can infect numerous lepidopteran hosts, primarily in the Noctuoidea, but most infections were atypical. Additionally, field collections in Bulgaria suggested that sympatric lepidopteran species do not serve as reservoirs for V. disparis (Solter et al. 2000). Taxonomic Summary
Fig. 55. Drawing from Timofejeva (1956), depicting octospores and individual spores of Thelohania disparis isolated from Lymantria dispar. The first eight stages pictured are developing octospores as they appear under light microscopy. The final drawing of spindle-shaped spores, considered by Timofejeva (1956) to be octospores released from the SV, probably represents diplokaryotic spores.
Vairimorpha disparis matures primarily in the fat body tissues, but gonads are also infected. In late stage infections, mature spores are sometimes found in isolated cells of the midgut epithelium and Malpighian tubules. Although primary spores and
Vairimorpha disparis (Timofejeva 1956) n. comb. Type host. Gypsy moth, Lymantria dispar (L.). Synonomy. Thelohania disparis Timofejeva (1956); Nosema muscularis Weiser (1957); Vairimorpha lymantriae (used provisionally), Nosema lymantriae1Thelohania similis Weiser (1957). Transmission. Per os. Although mature diplokaryotic spores and octospores form in the gonads, no conclusive evidence for transovum transmission is available. Site of infection and development in L. disparis larvae. The first merogonial cycle and primary sporulation (S1), before the secondary cycles (S2ss and S2oct), typically occur in the cytoplasm of midgut epithelial cells, midgut longitudinal and
Table 2. Lymantria dispar Vairimorpha sp. isolates held in liquid nitrogen storage at the Illinois Natural History Survey. Acc. no.a
GenBank no.
Sequence
Nosema sp. [Levishte]
Levishte, Bulgaria
1997-F
AF141129
ssrDNA
Vairimorpha sp. Vairimorpha sp. 3
Czech Rep. Czech Rep.
1985-B 1985-D
AF33315 L28974
ssrDNAb lsrDNA
Vairimorpha Vairimorpha Vairimorpha Vairimorpha Vairimorpha Vairimorpha
Czech Rep. Rupite Bulgaria Rimavska-Sobota, Slovakia Rupite, Bulgaria Rupite, Bulgaria Rupite, Bulgaria
1985-B 1985-C 1995-B 1995-C 1995-D 2004-A
L28976 L28970 Not entered Not entered DQ272237 Not entered
lsrDNAc lsrDNAc ssrDNAb ssrDNAb ssrDNAb ssrDNAb
Identity
a
Origin
sp. 4 sp. 5 sp. sp. sp. sp.
Number corresponds to year of collection. Identical ssrDNA sequences. c Identical lsrDNA sequences. b
Remarks/References Entered in GenBank as Vairimorpha lymantriae Solter et al. (2000) Maddox et al. (1999) Possibly a Nosema sp. Baker et al. (1994) Baker et al. (1994) Baker et al. (1994) Solter et al. (2000) This paper
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VAVRA ET AL.—VAIRIMORPHA DISPARIS N. COMB. IN LYMANTRIA DISPAR Table 3. Described Vairimorpha species from Lepidoptera. Identification
Host
Synonyms
Morphological Characters
Remarks/References
Vairimorpha necatrix
Pseudaletia unipuncta (Noctuidae)
Nosema necatrix Thelohania diazoma
S2ss: 12–15 pf coils S2oct: 12–14 pf coils Lamellar SV secretory products SSU rDNA distinct (Y00266a) LSU rDNA distinct (L28975)
Maddox and Sprenkel (1978), Mitchell and Cali (1993), Moore and Brooks (1992, 1994), Pilley (1976)
Vairimorpha ephestiae
Ephestia elutella (Pyralidae)
Thelohania ephestiae Octosporea ephestiae
S2ss: 14–16 pf coils Lamellar SV secretory products
Hazard and Oldacre (1975), Mattes (1928), Weiser (1961), Weiser and Purrini (1985)
Vairimorpha heterosporum
Plodia interpunctella (Pyralidae)
Nosema heterosporum
LSU rDNA distinct (L28973)
Baker et al. (1994), Kellen and Lindegren (1969)
Vairimorpha plodiae
Plodia interpunctella (Pyralidae)
Nosema plodiae
Lamellar SV secretory products
Kellen and Lindegren (1968), Malone and Canning (1982)
S2oct stage is abortive SSU rDNA distinct (AJ131645)
Canning et al. (1999)
Vairimorpha imperfecta Plutella xylostella (L.) (Plutellidae) Vairimorpha mesnili
Pieris brassicae, Pieris rapae (Pieridae)
Perezia mesnili Nosema mesnili Glugea mesnili Thelohania mesnlli
S2ss: 10–13 pf coils Malone and McIvor (1996), SSU rDNA distinct (no GenBank Paillot (1918) entry by Malone and McIvor 1996)
Vairimorpha antheraeae Antherea pernyi (Saturniidae)
Nosema antheraeae
Not infective to L. dispar
Simchuk, Lysenko, and Tchetkarova (1979), Yefimenko, Sokolova, and Issi (1990)
Vairimorpha disparis
Thelohania disparis Nosema muscularis Nosema lymantriae Thelohania similis
S2ss: 11–13 pf coils S2oct: 30 pf coils Mesh-like SV secretory products SSU rDNA distinct (DQ272237) LSU rDNA distinct (L28970)
Baker et al. (1994), Timofejeva (1956)
Lymantria dispar (Lymantriidae)
a GenBank accession number. pf, polar filament; SV, sporophorous vesicle.
transverse muscle cells, and connective tissue cells around the gut, as well as in the fat body. The secondary (S2ss and S2oct) merogony cycles typically occur in the fat body tissues where both diplokaryotic single spores and monokaryotic octospores are formed. Both S1 and S2 cycles can occur in other tissues, particularly in heavy, late-stage infections, but the number of spores formed is severely limited and, in some tissues, the spores do not mature. Spores. Three types of spores: (1) internally infective diplokaryotic primary spores with a conspicuous posterior vacuole, (elongate ellipsoidal, 5.4 2.5 mm, fresh); (2) secondary spores for inter-host transmission—diplokaryotic single spores (elongate ellipsoidal, 5.1 2.6 mm, fresh); and (3) secondary spores for inter-host transmission—monokaryotic octospores (broadly oval, 4.6 2.8 mm, fresh) enclosed in a semipersistent SV (7.3– 10.5 mm). Polar filament coils in all spore types are of the isofilar type: the primary spores have 4–5 (rarely 6) coils; the secondary single spores have 11–13 coils; and the octospores approximately 30 coils. Type locality. The material for this re-description was isolated from Lymantria dispar larvae collected from Salix alba in Rupite, Bulgaria (23116 0 1100 East; 41128 0 5600 North) in May 1995. Deposition of type specimens. Living spores of this isolate (Accession no. 1995-D) are maintained in the liquid nitrogen collections at the Illinois Natural History Survey, at Charles University, Prague, Czech Republic, and at BOKU—University
of Natural Resources and Applied Life Sciences, Vienna, Austria. In addition, Giemsa-stained slides are held at the Smithsonian Institute, Washington, DC, USA (Accession no. USNM 1084299), and in the collections of L. Solter, Illinois Natural History Survey, and J. Vavra and J. Weiser, Institute of Entomology, Academy of Sciences, Charles University, Czech Republic. The nucleotide sequence of the SSU and ITS rDNA gene is deposited in NCBI GenBank, Accession no. DQ272237.
ACKNOWLEDGMENTS The authors thank Mrs. Eva Kirchmanova, Charles University, Prague for expert assistance with electron microscopy, and gratefully acknowledge support from their institutes: Charles University and University of South Bohemia, Czech Republic; Illinois Natural History Survey; Connecticut Agricultural Experiment Station; Bulgarian Academy of Sciences; Fachhochschule Eberswalde; BOKU—University of Natural Resources and Applied Life Sciences, Vienna; USDA Forest Service, and funding from Agricultural Experiment Station Project no. ILLU-65-0344, USDA FS Cooperative Agreement no. AG 01CA-11242343-107, Charles University/USDA Forest Service Cooperative Agreement no. 161/79-982111; Czech Acad. of Sci., Project no. Z60220518; Charles University Grant no. MSM0021620828; and DFG—Deutsche Forschungsgemeinschaft no. 436 BUL 17/8/04.
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