Chicken as a Developmental Model

Chicken as a Developmental Model

Introductory article Article Contents • Introduction

Gi F Mok, University of East Anglia, Norwich, UK Abdulmajeed F Alrefaei, University of East Anglia, Norwich, UK James McColl, University of East Anglia, Norwich, UK Tim Grocott, University of East Anglia, Norwich, UK Andrea Münsterberg, University of East Anglia, Norwich, UK

• Stages of Chick Embryo Development • Manipulation of the Embryo – Classic Experiments • The Molecular Era: Chromosomes and Genome Information • Approaches to Study Gene Function • Transgenic Chickens • Embryo Culture and Live Imaging • The Chicken Model in Biomedical Research • Conclusion

Online posting date: 27th January 2015

The development of a complex organism from a single cell, the fertilised egg, has fascinated people for centuries. Embryo development is highly reproducible and exquisitely regulated. How is it that all tissues and organs form in the right places and at the right time? How is the development of different organ systems coordinated, so that they all fit together correctly at the end? It is challenging to study development, because many embryos are small or inaccessible. The chick embryo is a popular model system with many experimental advantages, which include classic ‘cut and paste’ experiments and mechanistic gene function analyses. The combination of micromanipulations with gain- or loss-of-function is particularly powerful. The recent development of transgenic lines and advanced imaging techniques ensure that the chicken remains an attractive model system, which will continue to make major contributions to our understanding of molecular and cellular mechanisms controlling developmental processes.

Introduction The chick embryo is a classic model to study development in a higher vertebrate, an amniote species. One reason for its popularity is the fact that embryos are easily accessible because they eLS subject area: Developmental Biology How to cite: Mok, Gi F; Alrefaei, Abdulmajeed F; McColl, James; Grocott, Tim; and Münsterberg, Andrea (January 2015) Chicken as a Developmental Model. In: eLS. John Wiley & Sons, Ltd: Chichester. DOI: 10.1002/9780470015902.a0021543

develop almost entirely outside the mother. By cutting a ‘window’ into the egg shell, the embryo can be revealed and its growth observed under a low-power microscope (Stern, 2005; Korn and Cramer, 2007). Following manipulations, such as microsurgery or introduction of foreign genetic material, drugs or viruses, the egg can be resealed and re-incubated. When the desired stage of development is reached, embryos can be recovered and harvested from the egg. The consequences of the manipulation are then investigated using a number of techniques, including cellular, molecular and biochemical methods. The chicken genome has been sequenced and molecular and genetic tools are available to interfere with gene function. Through these approaches, work in chick embryos has made important contributions to the elucidation of major concepts in vertebrate development, such as the concept of positional information (see also: Positional Information). Modern molecular tools applied in chick embryos enable us to decipher the mechanisms underlying developmental conditions and malformations, including those observed in human.

Stages of Chick Embryo Development To work with chick embryos in the laboratory, it is usually not necessary to house and breed adult animals. Fertilised eggs can be obtained from poultry farms and following delivery, eggs are stored in cooled incubators at 16 ∘ C; for optimal results, eggs should only be stored for 1–2 weeks. When they are needed, eggs are incubated at 37–39 ∘ C for the number of hours or days required to reach the stage of development being studied. This makes working with chick embryos very convenient. The early stages of chick embryo development progress faster when compared to the mouse for example. The mouse is the most commonly used mammalian model species (see also: Mice as Experimental Organisms). However, chicks develop more slowly than the commonly used non-amniote vertebrate model species: the amphibian Xenopus laevis (see also: Xenopus as an Experimental Organism) and the zebrafish Danio rerio (see also: Zebrafish as an Experimental Organism). Chicks hatch

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Chicken as a Developmental Model

from the egg after approximately 21 days of incubation. The length of the gestation period can vary slightly, depending on the breed and storage/incubation conditions. When the egg is laid, it already contains a pre-gastrula-stage embryo that consists of a concentric disc composed of two layers of cells, the so-called blastoderm. The earliest steps of development, the first cell divisions or cleavage divisions, have been studied in chick embryos (Bellairs et al., 1978), although this is challenging as cleavage occurs while the egg travels along the oviduct. Despite the difficulty, it is possible to recover fertilised chicken eggs from the hen by gentle squeezing and a series of stages has been described for chick embryo development before laying (Eyal-Giladi and Kochav, 1976). In vertebrates, pre-gastrula development is easier to investigate in non-amniotes such as the frog, X. laevis, or the zebrafish, D. rerio. Even in mouse embryos, it is possible to perform in vitro fertilisation and to observe cleavage divisions ex vivo, in a culture dish. In chickens, most research is conducted on embryos post-laying. Hamburger and Hamilton defined these stages of chick embryo development through to hatching (HH stage 1 to 46), based on a number of structural hallmarks, such as the number of somites or the size and shape of limb buds. This staging system is independent of the length of incubation or incubation conditions and Hamburger–Hamilton (HH) stages form the reference standard for research laboratories around the world (Hamburger and Hamilton, 1951). An important early process in development is gastrulation, during which embryos undergo complex morphogenesis and transform from a two-layered disc, consisting of the epiblast (top) and hypoblast (bottom) layers, into a three-dimensional embryo, with clearly defined axes and recognisable features (Chuai et al., 2012). The three germ layers, ectoderm, mesoderm and endoderm, are established during gastrulation, when some epiblast cells undergo an epithelial to mesenchymal transition (EMT). These cells then ingress through the primitive streak (PS), a transient structure that marks the head to tail axis, and migrate towards their target territory. A refined stage series has been provided, which uses gene expression patterns and morphological features to characterise these PS stages in more detail (Lopez-Sanchez et al., 2005). Gastrulation is followed by neurulation, which involves the formation of the neural plate, the rolling up of the neural plate to form the neural tube, followed by its differentiation into brain and spinal cord (Schoenwolf, 1991). At the same time, the cardiac mesoderm generates the first functioning organ, the heart, which initially forms as a simple tube that starts beating after around 40 h of incubation. The so-called paraxial mesoderm flanking the neural tube generates somites, paired aggregates of cells, which form in a regular, repetitive sequence and which give the body its segmented nature (Pourquie, 2004). Somites contribute to the vertebral column and associated muscles and tendons (Christ et al., 2004; Scaal and Christ, 2004). During the third day of development, the limbs begin to form, first the prospective wing and then the leg. Both are visible initially as thickened ridges, which grow out into buds that lengthen and change shape, with cells differentiating over the next few days (Tickle, 1995). The discrete anatomical parts of the limb become apparent, skeletal muscle cells migrate into the bud from the somites, digits separate through the programmed death of 2

inter-digital cells, cartilage condensations form the scaffolds for the future bones, nerve axons arrive and blood vessels form, all in the correct place. All major organ systems are well-developed half-way through gestation; therefore, the majority of experimental investigations focus on stages before embryonic day 10.

Manipulation of the Embryo – Classic Experiments The major advantage provided by avian embryos, mainly from chicken and the related quail (Ainsworth et al., 2010), is the ease with which they can be surgically manipulated in ovo. With the aid of a needle made from sharpened tungsten wire, small groups of cells or tissues can be removed/ablated, or transplanted from a donor embryo into another host embryo, and the effects of this manipulation on development can be studied. Embryonic tissues can also be explanted and cultured ex vivo; the capacity to differentiate in isolation can then be examined using various molecular markers. One example of this is the study of signalling cross-talk during the development and differentiation of the eye and in particular the lens (Grocott et al., 2011). The removal or ablation of tissues is often complemented by transplantation or grafting of the tissue and these types of experiments helped elucidate the importance of ‘organisers’ or signalling centres, which influence or ‘instruct’ other neighbouring tissues in their development. Transplants can be grafted into an ectopic or ‘out-of-place’ position within a host embryo; this is called heterotypic and will likely disrupt normal development. For example, when Hensen’s node, a group of cells at the tip of the PS, was grafted ectopically into an early chick embryo, a secondary body axis formed. This identified Hensen’s node as the amniote organiser, the functional equivalent of the Spemann–Mangold organiser, which had previously been discovered in amphibians (see also: Xenopus as an Experimental Organism; Stern, 2005). Similarly, the transplantation of a piece of notochord, a rod of mesoderm cells that normally resides below the neural tube, to an ectopic position on the opposite side resulted in patterning defects of the developing neural tube along the dorso-ventral axis (Yamada et al., 1991). This identified the notochord, a transient embryonic structure, as another important signalling centre. Other classic examples are transplantation experiments, which defined the zone of polarising activity (ZPA) and the apical ectodermal ridge (AER) as crucial regions that govern the appropriate development of vertebrate limbs (Tickle, 2004). In both of these examples, the transplanted tissue led to the induction of an ectopic structure. The ectopic notochord induced an ectopic floor plate, and the ectopic polarising region induced mirror image duplication of digits (Yamada et al., 1991; Tickle, 2004). In order to determine which signals or growth factors mimic the activity of the grafted tissue or cells, bead experiments can be performed (Figure 1). For the two examples mentioned, a bead soaked in recombinant Sonic hedgehog protein (Shh), a secreted peptide growth factor, would have the same effect as the grafted notochord or polarising region. Similar approaches can be used to revert or ‘rescue’ the effect of tissue ablation. For example,

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Chicken as a Developmental Model

AER so ht

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Figure 1 Bead implantation and detection of gene expression patterns by in situ hybridisation or immunostaining. (a) Beads soaked in growth factors or pharmacological inhibitors or activators of signalling pathways can be implanted into developing embryos. A bead implanted into the forelimb is shown. (b) Detection of specific messenger RNA transcripts in whole-mount embryos with anti-sense RNA probes against Fgf-8 (b), Myogenin = Mgn (c), or ventricular myosin heavy chain = vMHC (d). Probes incorporate a DIG-UTP nucleotide, which is detected using an alkaline phosphatase-coupled anti-DIG antibody. Alkaline phosphatase enzyme converts a substrate into a coloured precipitate, which generates a localised signal that is easily detected. Different structures are indicated by arrows in the different panels, AER = apical ectodermal ridge of the limb bud, so = somite, ht = heart. Other structures also expressing these genes are not indicated. (e) Protein can be detected by antibody staining in whole mount; the fluorescent signal indicates localisation of vMHC.

removal of the AER of a developing limb bud leads to stunted limb growth and this can be rescued with the application of a bead that is soaked in fibroblast growth factor (FGF). These types of experiment have been instrumental in identifying developmental signals involved in tissue patterning and cell fate determination. With the availability of specific pharmacological inhibitors of signal transduction cascades, it is possible to dissect the events downstream of receptor activation by growth factors. These small compound inhibitors can, for example, block phosphorylation events. They are applied locally by loading them on synthetic beads, which are implanted into the embryo close to the target tissue (Eblaghie et al., 2003). The exposure of developing tissues to synthetic agonists or antagonists of developmental signalling pathways, both in vivo and in explant culture, has helped elucidate signalling cross-talk and gene regulatory networks (Grocott et al., 2011; Streit et al., 2013). The chick embryo has also been used extensively for fate mapping studies. This involves the transplantation of tissues from a donor into an equivalent position within a host. If the cells in such a homotypic graft are labelled, they can be detected and therefore followed during development and thus, their normal ‘fate’ can be established. Classic approaches used quail-chick chimeras, in which quail tissue was grafted into a chick host (Le Douarin et al., 2008). Quail cells can be detected within the chicken using a specific nuclear stain or a quail-specific antibody. Other approaches to map the fate and developmental potential of groups of cells include labelling cells using the injection of dyes that are fluorescent. A disadvantage of this method is that the dye gets diluted over time, as cells divide, and the fluorescent signal may become too weak and too difficult to detect. Alternatives include the labelling of cells or tissue grafts by transfection, or more recently the use of transgenic embryos as donors (see later). For labelling by transfection, expression plasmids encoding green fluorescent protein (GFP) can be introduced by targeted microinjection and electroporation (see later).

The Molecular Era: Chromosomes and Genome Information Many avian species, including the chicken, have a large number of small chromosomes. In the chicken, it has only recently become possible to distinguish mini-chromosomes by karyotype analysis using chromosome paints (Griffin et al., 1999; Masabanda et al., 2004). In birds, females are the heterogametic sex and they have a female specific W-chromosome along with a Z-chromosome, males are the homogametic sex and have two Z-chromosomes. The mechanisms underlying sex determination are still not completely understood, although candidate sex determining genes have been identified (Smith et al., 2009). In addition, mixed-sex chimeras generated through tissue transplantation have demonstrated that there is a component determining somatic sex, which is cell autonomous; this has been termed cell autonomous sex identity (CASI) (Clinton et al., 2012). A draft genome of the red jungle fowl was produced in 2004 and it has helped annotate mammalian genomes through comparative genomics. In addition, the chicken genome has provided insights about genome evolution (Consortium, 2004; Wallis et al., 2004). Other molecular resources include a large collection of expressed sequence tags (ESTs) identified in a number of different tissues and stages of development (Hubbard et al., 2005). This has greatly facilitated the generation of molecular tools, for example, the preparation of anti-sense RNA probes to detect gene expression patterns by whole-mount in situ hybridisation (Figure 1). In addition to the detection of messenger RNA transcripts, it is also possible to detect proteins by whole-mount immunofluorescence using antibodies. Many chick-specific antibodies are available from the ‘Developmental Studies Hybridoma Bank’ (DSHB) at the University of Iowa. Antibodies, which were raised against proteins from human or mouse, also often cross-react with the chick homolog.

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ey

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Figure 2 Microinjection and electroporation of chicken embryos in EC culture or in ovo. (a) The injection set-up consists of a stereo-dissection microscope, micromanipulators, a pressure injector, an electroporator and a light source. (b) Close-up of the EC-culture dish, which contains the embryo mounted on a filter paper frame and placed on a semi-solid medium. (c) Close-up of the HH3 embryo in the filter paper carrier, illustrating the placement of the electrodes on either side of the primitive streak, indicated by a stippled line. (d) An embryo, which was electroporated at HH3 with a GFP-plasmid and incubated to HH5. The GFP-expressing cells have migrated away from the site of injection (bright green) and are now distributed in an arc shape. Many of these cells are prospective cardiac cells and will migrate to form the heart. (e) Electroporation of an embryo in ovo. Black ink injected beneath the embryo helps visualise it. (f) A chick embryo specifically expressing GFP in one half of the neural tube (nt), other structures indicated are the eye (ey) and the heart (ht), which is filled with red blood cells. (g) Section through the embryo shown in (f) shows restricted expression of the electroporated GFP plasmid on one side of the neural tube (nt).

Approaches to Study Gene Function The availability of genome sequence and ESTs has made the analysis of gene function more straightforward. Expression vectors can be constructed quickly using PCR-based cloning methods, and using microinjection in combination with electroporation, the DNA is introduced into different target tissues (Itasaki et al., 1999; Scaal et al., 2004; Voiculescu et al., 2008). Often, GFP is expressed from the same plasmid vector and thus the cells that have been successfully transfected are easily identified. Electroporation involves the application of repeated short pulses of an electrical current to the embryo. This transiently disrupts the plasma membrane, creating pores, which facilitate the uptake of DNA and other charged molecules by embryonic cells, and can be performed either in ovo or in embryo culture (ex ovo) (Figure 2). Approaches include the overexpression of genes of interest in order to examine the effects on cell fate, cell differentiation or tissue development. For example, the ectopic expression of a class of muscle-specific transcription factors in the developing neural tube leads to ectopic expression of muscle differentiation markers in neurons, thus confirming these transcription factors as master regulators of muscle fate (Sweetman et al., 2008). Furthermore, inducible gene expression systems have been developed, whereby application of tetracycline switches a promoter on or off (Watanabe et al., 2007). This allows investigation of stage-specific effects of a gene of interest and is useful if overexpression of this gene causes fatal consequences at earlier time points. In order to study longer lasting effects of a transgene, it is also possible to stably integrate vectors into the chicken genome using transposon-mediated gene transfer in combination with electroporation (Sato et al., 2007). Induction of a stably integrated transgene by administration of tetracycline enables the study of developmental processes at relatively late stages, for example, at embryonic days E6 to E8, where organ remodelling and differentiation is underway. In addition to microinjection and electroporation, targeted mis-expression of genes can also be mediated using an avian-specific retrovirus (RCAS, replication competent avian 4

sarcoma virus) as a shuttle vector (Morgan and Fekete, 1996). For example, it has been shown that infection of the neural tube with an RCAS virus expressing the paired-box transcription factor Pax3 leads to activation of skeletal-muscle-specific genes in this tissue (Maroto et al., 1997). This identified Pax3 as a key regulator for skeletal muscle development and differentiation. RCAS or plasmid vectors can both be used to express full-length coding regions, deletion mutants, tagged proteins or proteins that have been engineered to be constitutively active or dominantly negative (Abu-Elmagd et al., 2010). Similarly, RCAS or plasmids can be used to deliver short hairpin constructs, which use RNA interference (RNAi) to knock-down gene function (Harpavat and Cepko, 2006). Gene knock-down or loss-of-function can also be achieved by directly injecting and electroporating morpholino oligos (MO), which interfere with translation or splicing of messenger RNA (Norris and Streit, 2014). The MOs are often labelled with a covalently bound fluorescent dye (e.g. FITC, fluorescein isothiocyanate), which helps locate them. In addition, gene silencing can be achieved by directly injecting and electroporating double-stranded (ds)RNA, which also acts through RNAi. This is particularly efficient in the neural tube (Pekarik et al., 2003). Furthermore, anti-sense oligos can be delivered to developing limb buds or heart using pluronic gel (Becker et al., 1999; Rutland et al., 2009). A specific case of gene knock-down is the inhibition of short, non-coding RNAs, so-called microRNAs, using antagomir oligos (Goljanek-Whysall et al., 2014). Antagomirs can simply be injected into the target tissue, they do not require electroporation as they are modified with a cholesterol fatty acid, which facilitates their uptake into cells across the lipid bilayer.

Transgenic Chickens The methods described earlier use transient transgenesis and allow efficient alteration of gene function during the early stages of embryonic development. One disadvantage of using the chicken model is that genetic analysis is difficult due to late sexual maturity and the long generation time. However, methods

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Chicken as a Developmental Model

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Figure 3 Fate mapping using GFP-transgenic chicken embryos. (a) A single somite, micro-dissected from a GFP-transgenic embryo, was grafted into a non-transgenic host embryo (white arrow in b). (c) After 7 days of incubation, the somite has contributed to the vertebral column in the neck region, as indicated by the presence of GFP-positive cells. (d) More detailed analysis shows that a single somite contributes cells (purple colour) to two different vertebrae, which are indicated by a stippled outline. The phenomenon, whereby the original segments (=somites) contribute to the posterior and anterior half of new segments (=vertebrae) is called ‘re-segmentation’. (Courtesy of Mike McGrew, Roslin Institute, Edinburgh.)

CSF1R-mApple macrophage reporter (a)

Limb bud (b)

Limb bud (c)

Skeletal muscle (d)

Figure 4 Transgenic chick embryos expressing fluorescent markers, mApple (red) or eGFP (green), specifically in macrophages, under the control of the CSF1R promoter. (a) Macrophages (red) are distributed throughout the developing embryo. (b,c) Macrophages in limbs (green) at early (b) or later (c) stages of development. In (c), the limb has been co-stained with Lysotracker, a chemical that is picked up by active macrophages that are cleaning up apoptotic cells between the forming digits. These macrophages therefore appear yellow. (d) Distribution of macrophages (red) in skeletal muscle tissue. (Courtesy of Adam Balic, David Hume and Helen Sang, Roslin Institute, Edinburgh; see also Balic et al., 2014.)

have been developed to generate transgenic birds, chicken and quail, using lentiviral delivery of genetic material (McGrew et al., 2004; Chapman et al., 2005; Poynter et al., 2009). For example, transgenic chickens expressing GFP ubiquitously, in all cells and tissues, are an excellent source of labelled donor tissue to perform classic transplantation and fate mapping experiments (McGrew et al., 2008; Figure 3). The use of specific promoters makes it possible to restrict GFP expression to a subset of cells or a particular tissue, for example, to macrophages (Figure 4). This not only allows the detailed study of macrophage development but also enables studying the behaviour and migration of these cells during wound healing and regeneration (Balic et al., 2014). Similar studies in quail have allowed visualisation of blood vessel formation in live embryos using endothelial-specific promoters

controlling the expression of YFP (Sato et al., 2010). In future, it will be possible to generate transgenic reporter lines with the coding region for a fluorescent protein coupled to enhancers or promoters responsive to particular signalling pathways that are important during development, such as the Wnt or Notch pathways. Such transgenes would produce fluorescence only in the cells and tissues where they become activated, and if the fluorescence is short-lived, they would faithfully report pathway activity. Proof of principle for this approach has already been obtained using transient transgenesis, for example, by electroporation of Wnt reporters into developing somites (Rios et al., 2010). Genome editing approaches should in principle also be feasible in the chicken model; however, they are not yet routinely established.

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Embryo Culture and Live Imaging A number of culture methods exist to facilitate the study of embryo development ex ovo. One of the most popular methods for early embryos involves a filter paper ‘frame’, which is used to lift the embryo off the yolk. The filter paper frame serves as a carrier to transfer the embryo to a dish containing a layer of semi-solid medium made from egg white and agar (Chapman et al., 2001). This is useful to culture chick embryos for up to 2 days. For slightly longer culture periods, up to 3.5 days, the modified Cornish pasty method can be used (Nagai et al., 2014). In this method, the blastoderm is folded in half along the embryonic axis, the blastoderm edges are sealed, giving rise to a pouch or vesicle with the embryo sitting on top. Because the early chick embryo is flat and its normal development can be supported by suitable culture conditions, it has been possible to establish approaches to image developmental processes, such as cell migration (see also: Cell Migration during Development), in intact embryos in real time. Live imaging is usually combined with electroporation of GFP encoding plasmids in order to label specific populations of cells, which can then be followed (Chuai et al., 2009). For example, the movements of cells in pre-gastrula stage embryos have been extensively studied to reveal how the PS is formed (Chuai et al., 2006; Voiculescu et al., 2007) and the migration of prospective mesoderm cells has been investigated during gastrulation (Yang et al., 2002; Zamir et al., 2006; Iimura et al., 2007). It is possible to reveal the migration paths or trajectories of cells using image analysis programmes (Yang et al., 2002). When these approaches are combined with gain-of-function or dominant-negative interference, the molecular players and signals that are important in controlling cell behaviour can be dissected in the intact organism (Song et al., 2014; Yue et al., 2008). Similarly, it is possible to examine the contribution of the extracellular matrix (ECM) to cellular movements by labelling the ECM directly (Zamir et al., 2008). Further development of imaging techniques will enable the observation of cell migration and morphogenetic processes in larger embryos, and advanced image analysis methods will no doubt greatly enhance the information that can be obtained from this type of experiment.

The Chicken Model in Biomedical Research Many of the approaches described earlier have been instrumental for the investigation not only of normal development but also of the potential mechanisms that lead to human congenital malformations. When examining gene function in chick embryos, it often becomes apparent that gene knock-down or gain-of-function mimics aspects of human developmental defects, such as spina bifida, limb malformations, cleft palate or cardiac defects. Approaches using transient transgenesis in chick embryos can be complemented by genetic studies, for example, in mammalian (mouse) embryos, which use reverse genetics to completely remove (knock-out) gene function (see also: Mice as Experimental Organisms). The resulting phenotypes in chick 6

and mouse embryos are often similar as gene function tends to be conserved between closely related species. It is therefore also possible to extrapolate to human conditions and important mechanistic information can be obtained from these and other model organisms. This is nicely illustrated with examples from limb development (see also: Molecular Genetics of Human Congenital Limb Malformations). Many embryological or genetic manipulations affect limb outgrowth and/or the number and type of digits that form, thus providing insights into human limb congenital malformations, such as polydactyly (Sanz-Ezquerro and Tickle, 2003; Towers et al., 2008). Another example is the identification of the talpid3 gene in chick embryos (Davey et al., 2006); its functional characterisation in both chick and mouse (Bangs et al., 2011) has revealed the importance of primary cilia for the development of multiple organ systems, including craniofacial, neural tube, blood vessel and limb development, and has contributed to the classification of human ciliopathies, a group of conditions resulting from defects in primary cilia (Sharma et al., 2008). In addition, in ovo application of chemical compounds, such as vitamins, caffeine, alcohol or drugs have revealed potential teratogenic effects that they may have on the developing embryo, for example, the developing eye and nervous system (Flentke et al., 2014; Ma et al., 2014). In the chick limb, it was shown that thalidomide inhibits the formation of blood vessels in the growing bud, suggesting a possible mechanism by which this drug induced congenital limb defects in human, and indicating potential therapeutic applications inhibiting tumour angiogenesis (Therapontos et al., 2009). Additional examples of the chicken in biomedical research are covered in Further Reading.

Conclusion Chick embryos offer a number of advantages for experimental embryology and transient transgenesis. This is widely recognised and will ensure that the chicken model continues to make valuable contributions in modern bioscience research.

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Further Reading Brown WR, Hubbard SJ, Tickle C and Wilson SA (2003) The chicken as a model for large-scale analysis of vertebrate gene function. Nature Reviews Genetics 4 (2): 87–98. Davey MG and Tickle C (2007) The chicken as a model for embryonic development. Cytogenetic and Genome Research 117 (1–4): 231–239. Hilgers V, Pourquie O and Dubrulle J (2005) In vivo analysis of mRNA stability using the Tet-Off system in the chicken embryo. Developmental Biology 284 (2): 292–300. Hutson MR and Kirby ML (2007) Model systems for the study of heart development and disease. Cardiac neural crest and conotruncal malformations. Seminars in Cell & Developmental Biology 18 (1): 101–110. Kain KH, Miller JW, Jones-Paris CR, et al. (2014) The chick embryo as an expanding experimental model for cancer and cardiovascular research. Developmental Dynamics 243 (2): 216–228. Ogura T (2002) In vivo electroporation: a new frontier for gene delivery and embryology. Differentiation 70 (4–5): 163–171. Rashidi H and Sottile V (2009) The chick embryo: hatching a model for contemporary biomedical research. Bioessays 31 (4): 459–465.

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