Chicken as a Developmental Model

Chicken as aDevelopmental ModelGi 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

Introductory article

Article Contents• Introduction

• Stages of Chick Embryo Development

• Manipulation of the Embryo – ClassicExperiments

• The Molecular Era: Chromosomes and GenomeInformation

• Approaches to Study Gene Function

• Transgenic Chickens

• Embryo Culture and Live Imaging

• The Chicken Model in Biomedical Research

• Conclusion

th January 2015

The development of a complex organism from asingle cell, the fertilised egg, has fascinated peo-ple for centuries. Embryo development is highlyreproducible and exquisitely regulated. How is itthat all tissues and organs form in the right placesand at the right time? How is the development ofdifferent organ systems coordinated, so that theyall fit together correctly at the end? It is challeng-ing to study development, because many embryosare small or inaccessible. The chick embryo is apopular model system with many experimentaladvantages, which include classic ‘cut and paste’experiments and mechanistic gene function analy-ses. The combination of micromanipulations withgain- or loss-of-function is particularly power-ful. The recent development of transgenic linesand advanced imaging techniques ensure thatthe chicken remains an attractive model system,which will continue to make major contributionsto our understanding of molecular and cellularmechanisms controlling developmental processes.

Introduction

The chick embryo is a classic model to study development in ahigher vertebrate, an amniote species. One reason for its popu-larity 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 aDevelopmental Model. In: eLS. John Wiley & Sons, Ltd:Chichester.DOI: 10.1002/9780470015902.a0021543

develop almost entirely outside the mother. By cutting a ‘win-dow’ into the egg shell, the embryo can be revealed and its growthobserved under a low-power microscope (Stern, 2005; Korn andCramer, 2007). Following manipulations, such as microsurgeryor introduction of foreign genetic material, drugs or viruses, theegg can be resealed and re-incubated. When the desired stageof development is reached, embryos can be recovered and har-vested from the egg. The consequences of the manipulation arethen investigated using a number of techniques, including cel-lular, molecular and biochemical methods. The chicken genomehas been sequenced and molecular and genetic tools are availableto interfere with gene function. Through these approaches, workin chick embryos has made important contributions to the eluci-dation of major concepts in vertebrate development, such as theconcept of positional information (see also: Positional Informa-tion).Modernmolecular tools applied in chick embryos enable usto decipher themechanisms underlying developmental conditionsand malformations, including those observed in human.

Stages of Chick EmbryoDevelopment

To work with chick embryos in the laboratory, it is usually notnecessary to house and breed adult animals. Fertilised eggs canbe obtained from poultry farms and following delivery, eggs arestored in cooled incubators at 16 ∘C; for optimal results, eggsshould only be stored for 1–2 weeks. When they are needed,eggs are incubated at 37–39 ∘C for the number of hours or daysrequired to reach the stage of development being studied. Thismakes 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 themost commonly used mammalian model species (see also:Miceas Experimental Organisms). However, chicks develop moreslowly than the commonly used non-amniote vertebrate modelspecies: the amphibian Xenopus laevis (see also: Xenopus asan Experimental Organism) and the zebrafish Danio rerio (seealso: 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. Thelength of the gestation period can vary slightly, depending on thebreed and storage/incubation conditions. When the egg is laid,it already contains a pre-gastrula-stage embryo that consists ofa concentric disc composed of two layers of cells, the so-calledblastoderm. The earliest steps of development, the first cell divi-sions or cleavage divisions, have been studied in chick embryos(Bellairs et al., 1978), although this is challenging as cleavageoccurs while the egg travels along the oviduct. Despite the dif-ficulty, it is possible to recover fertilised chicken eggs from thehen by gentle squeezing and a series of stages has been describedfor chick embryo development before laying (Eyal-Giladi andKochav, 1976). In vertebrates, pre-gastrula development is eas-ier to investigate in non-amniotes such as the frog, X. laevis, orthe zebrafish, D. rerio. Even in mouse embryos, it is possible toperform in vitro fertilisation and to observe cleavage divisions exvivo, in a culture dish.In chickens, most research is conducted on embryos

post-laying. Hamburger and Hamilton defined these stagesof chick embryo development through to hatching (HH stage1 to 46), based on a number of structural hallmarks, such asthe number of somites or the size and shape of limb buds. Thisstaging system is independent of the length of incubation orincubation conditions and Hamburger–Hamilton (HH) stagesform the reference standard for research laboratories around theworld (Hamburger and Hamilton, 1951).An important early process in development is gastrulation, dur-

ing which embryos undergo complex morphogenesis and trans-form 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 endo-derm, are established during gastrulation, when some epiblastcells undergo an epithelial to mesenchymal transition (EMT).These cells then ingress through the primitive streak (PS), atransient structure that marks the head to tail axis, and migratetowards their target territory. A refined stage series has beenprovided, which uses gene expression patterns and morpho-logical features to characterise these PS stages in more detail(Lopez-Sanchez et al., 2005). Gastrulation is followed by neuru-lation, which involves the formation of the neural plate, the rollingup of the neural plate to form the neural tube, followed by its dif-ferentiation into brain and spinal cord (Schoenwolf, 1991). At thesame time, the cardiac mesoderm generates the first functioningorgan, the heart, which initially forms as a simple tube that startsbeating after around 40 h of incubation. The so-called parax-ial mesoderm flanking the neural tube generates somites, pairedaggregates of cells, which form in a regular, repetitive sequenceand which give the body its segmented nature (Pourquie, 2004).Somites contribute to the vertebral column and associated mus-cles 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 ini-tially as thickened ridges, which grow out into buds that lengthenand change shape, with cells differentiating over the next fewdays (Tickle, 1995). The discrete anatomical parts of the limbbecome apparent, skeletal muscle cells migrate into the bud fromthe somites, digits separate through the programmed death of

inter-digital cells, cartilage condensations form the scaffolds forthe future bones, nerve axons arrive and blood vessels form, allin the correct place. All major organ systems are well-developedhalf-way through gestation; therefore, the majority of experimen-tal investigations focus on stages before embryonic day 10.

Manipulation of theEmbryo – Classic Experiments

The major advantage provided by avian embryos, mainly fromchicken and the related quail (Ainsworth et al., 2010), is theease with which they can be surgically manipulated in ovo. Withthe aid of a needle made from sharpened tungsten wire, smallgroups of cells or tissues can be removed/ablated, or transplantedfrom a donor embryo into another host embryo, and the effectsof this manipulation on development can be studied. Embryonictissues can also be explanted and cultured ex vivo; the capacityto differentiate in isolation can then be examined using variousmolecular markers. One example of this is the study of signallingcross-talk during the development and differentiation of the eyeand 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 exper-iments helped elucidate the importance of ‘organisers’ or sig-nalling centres, which influence or ‘instruct’ other neighbour-ing tissues in their development. Transplants can be grafted intoan ectopic or ‘out-of-place’ position within a host embryo; thisis called heterotypic and will likely disrupt normal develop-ment. For example, when Hensen’s node, a group of cells atthe tip of the PS, was grafted ectopically into an early chickembryo, a secondary body axis formed. This identified Hensen’snode as the amniote organiser, the functional equivalent of theSpemann–Mangold organiser, which had previously been dis-covered in amphibians (see also: Xenopus as an ExperimentalOrganism; Stern, 2005).Similarly, the transplantation of a piece of notochord, a rod of

mesoderm cells that normally resides below the neural tube, toan ectopic position on the opposite side resulted in patterningdefects of the developing neural tube along the dorso-ventralaxis (Yamada et al., 1991). This identified the notochord, atransient embryonic structure, as another important signallingcentre. Other classic examples are transplantation experiments,which defined the zone of polarising activity (ZPA) and theapical ectodermal ridge (AER) as crucial regions that govern theappropriate development of vertebrate limbs (Tickle, 2004). Inboth of these examples, the transplanted tissue led to the inductionof an ectopic structure. The ectopic notochord induced an ectopicfloor plate, and the ectopic polarising region induced mirrorimage duplication of digits (Yamada et al., 1991; Tickle, 2004).In order to determine which signals or growth factors mimic theactivity of the grafted tissue or cells, bead experiments can beperformed (Figure 1). For the two examples mentioned, a beadsoaked in recombinant Sonic hedgehog protein (Shh), a secretedpeptide growth factor, would have the same effect as the graftednotochord or polarising region. Similar approaches can be usedto revert or ‘rescue’ the effect of tissue ablation. For example,

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(a) (b) (c) (d) (e)MgnFgf-8

Bead implant

vMHC

ht

soAER

vMHC

ht

Figure 1 Bead implantation and detection of gene expression patterns by in situ hybridisation or immunostaining. (a) Beads soakedin growth factors or pharmacological inhibitors or activators of signalling pathways can be implanted into developing embryos. A bead implantedinto 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 alkalinephosphatase-coupled anti-DIG antibody. Alkaline phosphatase enzyme converts a substrate into a coloured precipitate, which generates a localised signalthat 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 fluorescentsignal indicates localisation of vMHC.

removal of the AER of a developing limb bud leads to stuntedlimb growth and this can be rescued with the application of a beadthat is soaked in fibroblast growth factor (FGF). These types ofexperiment have been instrumental in identifying developmentalsignals involved in tissue patterning and cell fate determination.With the availability of specific pharmacological inhibitors ofsignal transduction cascades, it is possible to dissect the eventsdownstream of receptor activation by growth factors. These smallcompound inhibitors can, for example, block phosphorylationevents. They are applied locally by loading them on syntheticbeads, which are implanted into the embryo close to the targettissue (Eblaghie et al., 2003). The exposure of developing tissuesto synthetic agonists or antagonists of developmental signallingpathways, both in vivo and in explant culture, has helped elucidatesignalling 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 froma donor into an equivalent position within a host. If the cells insuch a homotypic graft are labelled, they can be detected andtherefore followed during development and thus, their normal‘fate’ can be established. Classic approaches used quail-chickchimeras, in which quail tissue was grafted into a chick host(Le Douarin et al., 2008). Quail cells can be detected withinthe chicken using a specific nuclear stain or a quail-specificantibody. Other approaches to map the fate and developmentalpotential of groups of cells include labelling cells using theinjection of dyes that are fluorescent. A disadvantage of thismethod is that the dye gets diluted over time, as cells divide,and the fluorescent signal may become too weak and too difficultto detect. Alternatives include the labelling of cells or tissuegrafts by transfection, or more recently the use of transgenicembryos 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: Chromosomesand Genome Information

Many avian species, including the chicken, have a large num-ber of small chromosomes. In the chicken, it has only recentlybecome possible to distinguish mini-chromosomes by karyotypeanalysis using chromosome paints (Griffin et al., 1999; Masa-banda et al., 2004). In birds, females are the heterogametic sexand they have a female specific W-chromosome along with aZ-chromosome, males are the homogametic sex and have twoZ-chromosomes. The mechanisms underlying sex determinationare still not completely understood, although candidate sex deter-mining genes have been identified (Smith et al., 2009). In addi-tion, mixed-sex chimeras generated through tissue transplanta-tion have demonstrated that there is a component determiningsomatic sex, which is cell autonomous; this has been termed cellautonomous 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 com-parative genomics. In addition, the chicken genome has providedinsights about genome evolution (Consortium, 2004;Wallis et al.,2004). Other molecular resources include a large collection ofexpressed sequence tags (ESTs) identified in a number of dif-ferent tissues and stages of development (Hubbard et al., 2005).This has greatly facilitated the generation of molecular tools, forexample, the preparation of anti-sense RNA probes to detect geneexpression patterns by whole-mount in situ hybridisation (Figure1). In addition to the detection of messenger RNA transcripts, itis also possible to detect proteins by whole-mount immunoflu-orescence using antibodies. Many chick-specific antibodies areavailable from the ‘Developmental Studies Hybridoma Bank’(DSHB) at the University of Iowa. Antibodies, which were raisedagainst proteins from human ormouse, also often cross-react withthe chick homolog.



(a) (b) (c) (d) (e) (f) (g)GFP GFP

Embryo

In ovo Section

Injection needle

Electrodes

EC culture dish

ntht

nt

ey

Figure 2 Microinjection and electroporation of chicken embryos in EC culture or in ovo. (a) The injection set-up consists of a stereo-dissectionmicroscope, micromanipulators, a pressure injector, an electroporator and a light source. (b) Close-up of the EC-culture dish, which contains the embryomounted 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 ofthe 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 andincubated 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 ofthese cells are prospective cardiac cells and will migrate to form the heart. (e) Electroporation of an embryo in ovo. Black ink injected beneath the embryohelps 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 oneside of the neural tube (nt).

Approaches to Study GeneFunction

The availability of genome sequence and ESTs hasmade the anal-ysis of gene function more straightforward. Expression vectorscan 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, GFPis expressed from the same plasmid vector and thus the cellsthat have been successfully transfected are easily identified. Elec-troporation involves the application of repeated short pulses ofan electrical current to the embryo. This transiently disrupts theplasma membrane, creating pores, which facilitate the uptake ofDNA and other charged molecules by embryonic cells, and canbe performed either in ovo or in embryo culture (ex ovo) (Figure2). Approaches include the overexpression of genes of interestin order to examine the effects on cell fate, cell differentiationor tissue development. For example, the ectopic expression of aclass of muscle-specific transcription factors in the developingneural tube leads to ectopic expression of muscle differentiationmarkers in neurons, thus confirming these transcription factorsas master regulators of muscle fate (Sweetman et al., 2008).Furthermore, inducible gene expression systems have been devel-oped, whereby application of tetracycline switches a promoteron or off (Watanabe et al., 2007). This allows investigation ofstage-specific effects of a gene of interest and is useful if overex-pression of this gene causes fatal consequences at earlier timepoints. In order to study longer lasting effects of a transgene,it is also possible to stably integrate vectors into the chickengenome using transposon-mediated gene transfer in combinationwith electroporation (Sato et al., 2007). Induction of a stably inte-grated transgene by administration of tetracycline enables thestudy of developmental processes at relatively late stages, forexample, at embryonic days E6 to E8, where organ remodellingand differentiation is underway.In addition to microinjection and electroporation, targeted

mis-expression of genes can also be mediated using anavian-specific retrovirus (RCAS, replication competent avian

sarcoma virus) as a shuttle vector (Morgan and Fekete, 1996).For example, it has been shown that infection of the neural tubewith an RCAS virus expressing the paired-box transcriptionfactor Pax3 leads to activation of skeletal-muscle-specific genesin this tissue (Maroto et al., 1997). This identified Pax3 as a keyregulator for skeletal muscle development and differentiation.RCAS or plasmid vectors can both be used to express full-lengthcoding regions, deletion mutants, tagged proteins or proteins thathave been engineered to be constitutively active or dominantlynegative (Abu-Elmagd et al., 2010). Similarly, RCAS or plasmidscan be used to deliver short hairpin constructs, which use RNAinterference (RNAi) to knock-down gene function (Harpavat andCepko, 2006). Gene knock-down or loss-of-function can also beachieved by directly injecting and electroporating morpholinooligos (MO), which interfere with translation or splicing ofmessenger RNA (Norris and Streit, 2014). The MOs are oftenlabelled 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 elec-troporating double-stranded (ds)RNA, which also acts throughRNAi. This is particularly efficient in the neural tube (Pekariket al., 2003). Furthermore, anti-sense oligos can be delivered todeveloping limb buds or heart using pluronic gel (Becker et al.,1999; Rutland et al., 2009). A specific case of gene knock-downis the inhibition of short, non-coding RNAs, so-called microR-NAs, using antagomir oligos (Goljanek-Whysall et al., 2014).Antagomirs can simply be injected into the target tissue, they donot require electroporation as they are modified with a choles-terol fatty acid, which facilitates their uptake into cells across thelipid bilayer.

Transgenic Chickens

The methods described earlier use transient transgenesis andallow efficient alteration of gene function during the early stagesof embryonic development. One disadvantage of using thechicken model is that genetic analysis is difficult due to latesexual maturity and the long generation time. However, methods



(d)

(c)(b)(a)

HH10 somite graft Vertebrae

HH32

Day 7Day 1

Figure 3 Fate mapping using GFP-transgenic chicken embryos. (a) A single somite, micro-dissected from a GFP-transgenic embryo, was graftedinto 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 differentvertebrae, which are indicated by a stippled outline. The phenomenon, whereby the original segments (=somites) contribute to the posterior and anteriorhalf of new segments (=vertebrae) is called ‘re-segmentation’. (Courtesy of Mike McGrew, Roslin Institute, Edinburgh.)

(a) (b)Limb bud

CSF1R-mApplemacrophage reporter Limb bud Skeletal muscle

(c) (d)

Figure 4 Transgenic chick embryos expressing fluorescent markers, mApple (red) or eGFP (green), specifically in macrophages, underthe control of the CSF1R promoter. (a) Macrophages (red) are distributed throughout the developing embryo. (b,c) Macrophages in limbs (green) atearly (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 thatare cleaning up apoptotic cells between the forming digits. These macrophages therefore appear yellow. (d) Distribution of macrophages (red) in skeletalmuscle 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 andquail, using lentiviral delivery of genetic material (McGrewet al., 2004; Chapman et al., 2005; Poynter et al., 2009). Forexample, transgenic chickens expressing GFP ubiquitously, in allcells and tissues, are an excellent source of labelled donor tissueto perform classic transplantation and fate mapping experiments(McGrew et al., 2008; Figure 3). The use of specific promotersmakes it possible to restrict GFP expression to a subset of cells ora particular tissue, for example, to macrophages (Figure 4). Thisnot only allows the detailed study of macrophage developmentbut also enables studying the behaviour and migration of thesecells during wound healing and regeneration (Balic et al., 2014).Similar studies in quail have allowed visualisation of blood vesselformation 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 thecoding region for a fluorescent protein coupled to enhancersor promoters responsive to particular signalling pathways thatare important during development, such as the Wnt or Notchpathways. Such transgenes would produce fluorescence only inthe cells and tissues where they become activated, and if thefluorescence is short-lived, they would faithfully report pathwayactivity. Proof of principle for this approach has already beenobtained using transient transgenesis, for example, by electro-poration of Wnt reporters into developing somites (Rios et al.,2010). Genome editing approaches should in principle also befeasible in the chicken model; however, they are not yet routinelyestablished.



Embryo Culture and Live Imaging

A number of culture methods exist to facilitate the study ofembryo development ex ovo. One of the most popular methodsfor early embryos involves a filter paper ‘frame’, which is usedto lift the embryo off the yolk. The filter paper frame serves asa carrier to transfer the embryo to a dish containing a layer ofsemi-solid medium made from egg white and agar (Chapmanet al., 2001). This is useful to culture chick embryos for upto 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 theembryonic axis, the blastoderm edges are sealed, giving rise to apouch or vesicle with the embryo sitting on top.Because the early chick embryo is flat and its normal devel-

opment can be supported by suitable culture conditions, it hasbeen possible to establish approaches to image developmentalprocesses, such as cell migration (see also: Cell Migration dur-ing Development), in intact embryos in real time. Live imagingis usually combined with electroporation of GFP encoding plas-mids in order to label specific populations of cells, which canthen be followed (Chuai et al., 2009). For example, the move-ments of cells in pre-gastrula stage embryos have been exten-sively studied to reveal how the PS is formed (Chuai et al., 2006;Voiculescu et al., 2007) and the migration of prospective meso-derm cells has been investigated during gastrulation (Yang et al.,2002; Zamir et al., 2006; Iimura et al., 2007). It is possible toreveal the migration paths or trajectories of cells using imageanalysis programmes (Yang et al., 2002). When these approachesare combined with gain-of-function or dominant-negative inter-ference, the molecular players and signals that are important incontrolling cell behaviour can be dissected in the intact organ-ism (Song et al., 2014; Yue et al., 2008). Similarly, it is possibleto examine the contribution of the extracellular matrix (ECM) tocellular movements by labelling the ECM directly (Zamir et al.,2008). Further development of imaging techniques will enablethe observation of cell migration and morphogenetic processesin larger embryos, and advanced image analysis methods will nodoubt greatly enhance the information that can be obtained fromthis type of experiment.

The Chicken Model in BiomedicalResearch

Many of the approaches described earlier have been instrumen-tal for the investigation not only of normal development butalso of the potential mechanisms that lead to human congen-ital malformations. When examining gene function in chickembryos, it often becomes apparent that gene knock-downor gain-of-function mimics aspects of human developmentaldefects, such as spina bifida, limb malformations, cleft palate orcardiac defects. Approaches using transient transgenesis in chickembryos can be complemented by genetic studies, for example,in mammalian (mouse) embryos, which use reverse genetics tocompletely remove (knock-out) gene function (see also:Mice asExperimental Organisms). The resulting phenotypes in chick

and mouse embryos are often similar as gene function tends tobe conserved between closely related species. It is therefore alsopossible to extrapolate to human conditions and important mech-anistic information can be obtained from these and other modelorganisms. This is nicely illustrated with examples from limbdevelopment (see also: Molecular Genetics of Human Con-genital Limb Malformations). Many embryological or geneticmanipulations affect limb outgrowth and/or the number and typeof digits that form, thus providing insights into human limbcongenital malformations, such as polydactyly (Sanz-Ezquerroand Tickle, 2003; Towers et al., 2008). Another example is theidentification 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 primarycilia for the development of multiple organ systems, includingcraniofacial, 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 potentialteratogenic effects that they may have on the developing embryo,for example, the developing eye and nervous system (Flentkeet al., 2014; Ma et al., 2014). In the chick limb, it was shownthat thalidomide inhibits the formation of blood vessels in thegrowing bud, suggesting a possible mechanism by which thisdrug induced congenital limb defects in human, and indicatingpotential therapeutic applications inhibiting tumour angiogenesis(Therapontos et al., 2009). Additional examples of the chicken inbiomedical research are covered in Further Reading.

Conclusion

Chick embryos offer a number of advantages for experimentalembryology and transient transgenesis. This is widely recognisedandwill ensure that the chickenmodel continues tomake valuablecontributions in modern bioscience research.

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Further Reading

BrownWR,Hubbard SJ, Tickle C andWilson SA (2003) The chickenas 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 embry-onic development. Cytogenetic and Genome Research 117 (1–4):231–239.

Hilgers V, Pourquie O and Dubrulle J (2005) In vivo analysis ofmRNA 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 ofheart development and disease. Cardiac neural crest and conotrun-cal malformations. Seminars in Cell & Developmental Biology 18(1): 101–110.

Kain KH, Miller JW, Jones-Paris CR, et al. (2014) The chick embryoas an expanding experimental model for cancer and cardiovascularresearch. Developmental Dynamics 243 (2): 216–228.

Ogura T (2002) In vivo electroporation: a new frontier for genedelivery and embryology. Differentiation 70 (4–5): 163–171.

Rashidi H and Sottile V (2009) The chick embryo: hatching a modelfor contemporary biomedical research. Bioessays 31 (4): 459–465.


Chicken as a Developmental Model

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