manipulating the avian egg: applications for embryo transfer, … · 2017-10-01 · avian biology...

AVIAN BIOLOGY RESEARCH 10 (3), 2017 146–155

Paper 1600746 https://doi.org/10.3184/175815617X14951979279268 www.avianbiologyresearch.co.uk

Manipulating the avian egg: applications for embryo transfer,

transgenics, and cloning

Michael E. Kjellanda,b,c*, Ben Novakd, Alice Blue-McLendone, Salvador Romof and

Duane C. Kraemerb

aEcological Systems Laboratory, Department of Wildlife & Fisheries Sciences, College of Agriculture

and Life Science, Texas A&M University, 210 Nagle Hall, College Station, TX 77843, USAbReproductive Sciences Laboratory, Department of Veterinary Physiology and Pharmacology, College of

Veterinary Medicine and Biomedical Sciences, Texas A&M University, College Station, TX 77843, USAcConservation, Genetics & Biotech, LLC, 10921 36 St. SE Valley City, ND 58072, USAdRevive & Restore, 323 Pine Street Suite D, Sausalito, CA 94965, USAeWinnie Carter Wildlife Center, Department of Veterinary Physiology and Pharmacology, College of

Veterinary Medicine and Biomedical Sciences, Texas A&M University, College Station, TX 77843, USA fFacultad de Estudios Superiores Cuautitlan, UNAM, Cuautitlan, Estado de Mexico, Mexico

*E-mail: [email protected]

ABSTRACT

In vitro production of germline chimeras and avian cloning may utilise the transfer of avian embryos from their original eggshell to a surrogate eggshell for culture during incubation. Such embryo transfer is valuable for avian cloning as the only alternative would be to transfer the cloned avian embryos into the infundibulum of recipient birds. Given the advances in paleogenomics, synthetic biology, and gene editing, a similar approach might be used to generate extinct species, i.e. de-extinction. One objective of the present research was to examine if ratite eggs could be manipulated via windowing and sham injection, similar to that which could allow for avian genome manipulation and subsequent development. The efficiency of interspecific avian embryo transfer using Chicken (Gallus gallus domesticus) donor eggs and Turkey (Meleagris gallopavo) recipient eggshells was also investigated. Egg windowing and embryo transfer techniques utilised in the present research were adapted from those found in the scientific literature. Presumed fertile eggs from Rhode Island Red (n = 40), Silkie (n = 2), and White Leghorn Chickens (n = 18), Turkey (n = 48), Emu (Dromaius novaehollandiae) (n = 79), and Ostrich (Struthio camelus) (n = 89) were used in this research. Of the 41 Chicken eggs used for transfers into recipient Turkey eggshells, only one (2.4%) produced a chick. Of 31 windowed Emu eggs, one embryo survived for 25 d but no chicks were produced. Of 36 windowed Ostrich eggs, one embryo survived and hatched. The efficiency of the windowing and embryo transfers to produce chicks was low and further refinements are needed. Importantly, the results herein establish that manipulating ratite embryos is possible.

Keywords: avian, Chicken, Gallus gallus domesticus, Turkey, Meleagris gallopavo, Emu, Dromaius novaehollandiae, Ostrich, Struthio camelus

1. INTRODUCTION

The ability to efficiently and precisely edit genomes with CRISPR/Cas9 has revolutionised genomic research (Pennisi, 2013; Barrangou, 2014). Targeted knock-ins, knock-outs, and precise base-pair editing have been reported with no off target events in a myriad of model organisms, and has recently expanded with genome editing of germlines and somatic cells in the Chicken, Gallus gallus domesticus (Véron et al., 2015; Dimitrov et al., 2016; Zuo et al., 2016). The ability to edit avian genomes in vitro and in ovo opens the door for applying genome engineering to avian conservation issues, such as

introducing genetic diversity to bottlenecked populations and facilitating genomic adaptation to introduced disease and climate change (Johnson et al., 2016).

Combined with advances in paleogenomics, precise genome editing in avian species establishes the platform to edit the genomes of living birds to express traits of extinct species for producing effective ecological replacements, a process dubbed as de-extinction (Shapiro, 2016a). The genomes of several extinct bird species including the Passenger Pigeon, Ectopistes migratorius (Shapiro et al., 2016), Heath Hen, Tympanuchus cupido cupido (Johnson et al., 2015), Dodo, Raphus cucullatus (Shapiro, 2016b), Great Auk, Pinguinus impennis (Gilbert, T., personal communication), and multiple Moa species (Dinornis

Manipulating the avian egg 147

sp.) (Coutier, A. and Iorns, D., personal communication) are already being mapped using extant species’ reference genomes. Among the aforementioned extinct species, active de-extinction projects are underway for the Passenger Pigeon and Heath Hen by the non-profit group Revive & Restore (www.reviverestore.org) and Moa de-extinction is being pursued by the non-profit group Genetic Rescue Foundation (www.geneticrescue.science). These projects are driving the expansion and development of advanced reproductive technologies for avian species with regard to producing genome edited offspring, of which there are two proven methods: (1) in ovo production of germline chimeras through direct injection of transfection agents to the subgerminal cavity (McGrew et al., 2004) or ventral dorsal aorta (Tyack et al., 2013); and (2) in vitro production of both intraspecific and interspecific germline chimeras with cultured primordial germ cells (PGCs) (van de Lavoir et al., 2006, 2012).

While no bird has yet been cloned through somatic cell nuclear transfer (SCNT), cloning could provide a means to produce genomically-edited offspring. According to Sang and Sherman (2003, 2005a,b,c), an avian embryo may be allowed to develop to term in vitro or in a host shell by a modification of the technique described in their avian cloning patent. According to the patent methodology, an egg is removed from a female bird, enucleated according to the method of the first aspect of the invention, fused with a donor nucleus to generate an embryo and allowed to develop to hatching within a host shell. Optionally, transfer of the contents of one egg to a larger egg could be conducted, to leave an air-space over the developing embryo. Whether or not this can be accomplished using cloned avian embryos remains to be determined; however, the first step – enucleation of the avian ovum – was successfully achieved in domestic Chickens recently (Kjelland et al., 2014), establishing the basis to pursue cloning in birds further.

Both established in vitro production of germline chimeras and proposed methods of avian cloning involve the transfer of avian embryos from their original eggshell to a different surrogate eggshell, which has been windowed

for incubation. Such embryo transfer can be used to rescue embryos from damaged eggshells, but is critical for avian cloning as the only alternative would be to transfer the cloned avian embryos into the infundibulum of recipient birds via surgery. Surgical ovum transfers add to the complexity of avian cloning experiments, compared to propagating the cloned embryos in a culture system of surrogate eggshells. Regardless of the approach used to generate de-extinct avian proxy species, embryo transfer may be required to hatch large flightless birds whose gigantism poses a severe problem for developing properly in the eggshells of their extant genomic template species (i.e. the species whose genomes will be edited to produce de-extinct proxies). It may be unlikely that appropriate interspecific germline chimeras can be produced to lay suitable sized eggs for the Great Auk, Pinguinus impennis (with a four fold larger egg than the genomic template species), the Dodo, Raphus cucullatus (eight fold larger egg), Elephant bird, Aepyornis maximus (20 fold larger egg), and Giant Moa, Dinornis robustus (64 fold larger egg) (see Table 1).

There is a clear need for a culture system to grow an avian embryo in a surrogate eggshell for embryo transfer, either an eggshell of the same species, an eggshell of a different species, or a synthetic eggshell, if genome editing is to reach its fullest potential for avian models. If interspecific embryo transfer in domestic fowl can be accomplished efficiently, this model could be a viable tool for the preservation of unique and rare strains of domestic fowl and endangered bird species in future conservation efforts when precious eggs become damaged in captive propagation programmes. However, as previously stated, determining the feasibility and perhaps improving the embryo culture system for use in cloning is imperative for avian cloning procedures. The cloning of birds as a tool for preservation and genetic rescue of unique and rare strains of domestic fowl and endangered avian species can be a viable tool in future conservation efforts, particularly for species in which PGCs for in vitro germline transmission are (1) difficult to procure (i.e. birds that are difficult to breed in captivity, are possibly extinct, or eggs cannot

Table 1 Calculated egg massesa of extinct and extant bird species

Extinct species Egg mass (g)Corresponding genomic template species

Egg mass (g) References

Elephant bird, Aepyornis maximus

7,762Kiwi,Apteryx sp.

378 Amadon, 1947; Mitchell et al., 2014

Moa,Dinornis sp.

4,008Tinamou,Tinamus major zuliensis

63 Amadon, 1947; Preston, 1968; Mitchell et al., 2014

Great Auk,Pinginis impennis

358Razorbill,Alca torda

89b Bengtson, 1984; Birkhead and Nettleship, 1984; Hipfner, 2000; Moum et al., 2002

Dodo,Raphus cucullatus

197c Nicobar Pigeon,Caloenas nicobarica

25 Livezey, 1993; Brown, 1995; Shapiro et al., 2002

aEgg masses listed are calculated as mass = volume = 0.5 LB2, where L is length and B is breadth of the egg per the assumption that mass and volume are approximately equal for freshly laid eggs (Amadon, 1947). bCalculated from an average of five mean volume indexes given by Birkhead and Nettleship (1984) and Hipfner (2000).cThis is a predicted value, as no egg of the Dodo bird was ever precisely measured, preserved for measurement, or has been discovered during paleontological excavations.

148 Michael E. Kjelland, Ben Novak, Alice Blue-McLendon, Salvador Romo and Duane C. Kraemer

feasibly be spared for PGC propagation) or (2) non-existent (which is the case for some extinct species, such as the Hawaiian Honeycreeper the Po’ouli, Melamprosops phaeosoma). Provided a Chicken could be cloned as a model avian species, the same technology could be used for the conservation of critically endangered bird species such as the Spix’s Macaw (Cyanopsitta spixii), which is represented by only about 60 individuals in the world (arguably eggs cannot be spared to expand the population through in vitro germline transmission). Provided avian cloning technology is eventually developed, it would further establish and facilitate the possibility of future reptilian cloning and de-extinction conservation efforts, in which a good candidate would be ‘Lonesome George’, the last member of one subspecies of Galapagos Tortoise (Chelonoidis nigra abingdonii) that went extinct in 2012 and from which cells are cryopreserved (Nicholls, 2012).

Currently, there is evidence of success with egg windowing, e.g. Perry (1988), and intra-species and inter-species embryo transfer using Chicken eggs and Turkey (Meleagris gallopavo) eggs, respectively, e.g. Borwornpinyo et al. (2005). However, no evidence can be found in the scientific literature regarding such manipulation using ratite eggs, which require different incubation conditions and a much longer incubation duration compared to domestic Chickens. The null hypothesis to be tested was that given different incubation requirements and egg morphologies for ratites, compared to domestic Chickens and domestic Turkeys, egg windowing with the subsequent hatching of chicks would not be possible. In order to test the aforementioned hypothesis, research regarding the possibility of ratite egg windowing and subsequent embryo viability and chick development was undertaken. More specifically, an objective of the present research was to examine if ratite eggs could be manipulated via windowing and with sham injection, similar to that which could allow for avian genome manipulation and subsequent development. We also investigated the efficiency of interspecific avian embryo transfer using Chicken donor eggs and recipient Turkey eggshells, as conducted in the past by Borwornpinyo et al. (2005). It should also be mentioned that other methods are being developed that use artificial materials instead of eggshells to grow chick embryos, e.g. Lai and Liu (2015).

2. METHODS

2.1 Chicken egg windowing and interspecific embryo transfer

Windowing procedures were modelled after Perry (1988), Rowlett and Simkiss (1987), and Borwornpinyo et al. (2005). All egg windowing procedures, except eggshell cap removal, were performed in a laminar flow hood.

Specifically, eggs were wiped with a 70% alcohol solution before handling. Eggshells were cut on the

blunt end containing the air chamber, without damage to the inner eggshell membrane, using a MultiPro® 2850-02 DREMEL tool (35,000 RPM) with a diamond wheel cutting disc (DREMEL® 545). The diamond wheel was also soaked in a 70% alcohol solution before use. The freshly cut eggshell edges were treated with 0.5 mg mL–1 of gentamicin. The cut eggshell piece was then removed using a number 10 surgical blade and then 0.2 mL of 0.5 mg mL–1 of gentamicin was added to the exposed albumin. Standard meat wrap, similar to cling wrap, was used as a windowing material to replace the removed eggshell piece and sealed with Elmer’s® glue. Other studies have also used Saran Wrap and Handi-Wrap® or egg thin albumin as a seal (Borwornpinyo et al., 2005). The egg yolks were then placed into recipient eggshells with albumin and windowed using the aforementioned procedure. Following embryo transfers, the modified eggs were placed into a Humidaire (Model 50) redwood drum-type incubator with automatic turner (45 degree rotation each way with cessation of turning on day 19) maintained at 37.5 °C and 60% relative humidity for approximately 21 days or until hatching.

Fertile eggs from Rhode Island Red (RIR, n = 40), Silkie (n = 2), and White Leghorn Chickens (n = 18) were obtained from the Poultry Science Center at Texas A&M University and Turkey eggs (n = 48) were obtained from the Southern Plains Agricultural Research Center, United States Department of Agriculture (USDA) and the Poultry Science Center at Texas A&M University. All Chicken eggs were less than 48 h old at the time of manipulation. Silkie (n = 2), White Leghorn (n = 12), and RIR (n = 27) eggs that were developing after two days of incubation were used for transfers into recipient Turkey eggshells (after removing the top portion of the eggshell, i.e. end with the natural air cell). The remaining Chicken eggs were used for substitute albumen as needed, e.g. if albumen loss occurred during the transfer to a recipient eggshell surrogate albumen was used to replace it.

2.2 Emu egg windowing and intraspecies embryo transfer

Fertile Emu eggs (n = 79) were collected from a free ranging Emu colony near Marquez, Texas. Eggs were weighed using a mass balance scale and the weights recorded. Length, width, and circumference measurements were also recorded. Of the total number of eggs, 24 were used as controls to assess fertility. Some eggs (n = 11) were windowed while others (n = 28) were used for transferring their contents to recipient eggshells followed by windowing. It was expected that some transfers would fail i.e. result in a broken yolk, and eggshells from those donor eggs were used as recipient eggshells when necessary, provided they were large enough for the donor egg contents. First, the air chamber was assumed to be located in the blunt end or uppermost portion of the egg as it lay horizontally, hereafter referred to as the blunt end-angle


method (BEAM) as described in Kjelland et al. (2012). Egg windowing procedures were the same as those described above with the exception that the smaller egg contents were transferred to larger recipient eggshells. The windowed and control eggs were placed into a GQF incubator (GQF Manufacturing Company Inc., Georgia, USA) and maintained at approximately 36.1–36.4 °C and 35% relative humidity until hatching (average of 49–52 days [Jefferey, 2001a]) or until loss of viability. Eggs were set with the windowed end up (i.e. air cell up) and turned through a 90 degree rotation four to six times per day as described in Jefferey (2001b). Studies have shown a 1–2% increase in the hatchability of eggs placed vertically with the air cell at the top of the egg over that of ratite eggs placed on their sides (Jeffrey et al., 1996; Kjelland et al., 2012).

2.3 Ostrich egg windowing and sham injection

Fertile Ostrich eggs (n = 89) were obtained from the Winnie Carter Wildlife Center at Texas A&M University. Eggs were weighed using a mass balance scale and the weights recorded. Of the total number of eggs, 36 were randomly selected for the windowing experiment using a Bernoulli distribution. Ostrich egg window sizes varied from approximately 6.35–7 cm in diameter. Two groups were formed: (1) eggs with the inner air cell membrane removed (n = 31) after windowing, i.e. windowing as described above but some with eggshell caps glued back on (Group 1); and (2) eggs with the inner air cell membrane left intact (n = 5) after windowing but sham injected, with or without the eggshell cap glued back on (Group 2). For the sham injection of Ostrich eggs a sterile glass micropipette for somatic cell nuclear transfer (SCNT) was used to pierce the egg air cell membrane and inserted into the egg cytoplasm at the point where the blastodisc would likely be located (i.e. simulating an injection without injecting anything into the egg). The sham injection was performed to simulate the

physical alteration of the membranes as would occur with PGC or transfection solution injections.

The air chamber was located by candling prior to windowing the egg. Egg candling is the technique of shining a light through an egg to observe embryo development or viability (Cartwright, 2000; Kjelland et al., 2012). The windowed eggs were placed into a NatureForm Nom 45 incubator (NatureForm Hatchery Technologies, Florida, USA) and maintained at 36–36.7 °C and 20–25% relative humidity until hatching (average of 42 days [Jefferey 2001b]) or loss of viability (inspected weekly). Eggs were set with the windowed end up (i.e. air cell up) and turned through a 90 degree rotation four to six times per day as described in Jefferey (2001b).

3. RESULTS

3.1 Chicken egg windowing and interspecific embryo transfer

Of the 41 Chicken eggs used for transfers into recipient Turkey eggshells, only one (2.4%) produced a chick. More specifically, of the RIR embryos (n = 27) transferred one chick was produced (Figure 1 and see ESI for videos), but none for the Silkie and White Leghorn windowed eggs. All but one embryo experienced mortality before stage 32 based on stages of chick development described by Hamburger and Hamilton (1951).

3.2 Emu egg windowing and intraspecies embryo transfer

The mean (μ) and standard deviation (SD), minimum (min), and maximum (max) for egg weight (g), length (mm), width (mm), and circumference (mm) were the following: μ = 577.51, SD = 42.917, min = 421.5, max = 659.3; μ =

Figure 1 (a) Rhode Island Red Chicken embryo in windowed egg, (b) Chicken embryo removed from recipient Turkey eggshell before hatching to assess yolk absorption, subsequently placed back into eggshell, (c) Chicken embryo in recipient Turkey eggshell several hours before hatching, standard meat wrap windowing material removed and replaced with a sterile Petri dish to cover embryo, (d) chick hatched from interspecific embryo transfer egg, i.e. Chicken embryo transferred to recipient Turkey eggshell and windowed, and (e) adult Chicken hatched from embryo transfer windowed egg (note: same individual in all photos).


130.74, SD = 4.198, min = 120.1, max = 142.8; μ = 87.74, SD = 2.01, min = 84, max = 92; and μ = 280.73, SD = 7.032, min = 268, max = 295, respectively (Figures 2 and 3). Of the control (i.e. non-windowed) eggs, 3 of 24 (12.5%) did not show any sign of development and were likely infertile. Of the 28 attempted donor egg content transfers to recipient

eggshells, 20 were transferred and windowed successfully, a 71.4% transfer success rate. The egg yolk broke in a small number of the egg content transfers (n = 8).

Of 31 windowed Emu eggs (i.e. windowed without transfer [n = 11]) and transfer windowed (n = 20), only one embryo developed and survived for 25 d (Figure 4) with the

Figure 3 Emu egg (n = 79) box and whisker plots for (a) weight (g), (b) length (mm), (c) width (mm), and (d) circumference (mm).

Figure 2 Emu egg (n = 79) characteristics: (a) weight (g); (b) length (mm); (c) width (mm); (d) circumference (mm); and (e) relationship between length, width, and circumference.


second longest survivor lasting 18 d; although no chicks were successfully hatched. Both of the longest surviving embryos were from egg content transfer windowed eggs. All other embryos experienced mortality earlier in development, i.e. on or before stage 28, based on chick embryo stages of development by Hamburger and Hamilton (1951). Emu embryos typically take 2–3 times the incubation time to reach equivalent Chicken stages (Nagai et al., 2011), i.e. stages described by Hamburger and Hamilton (1951).

3.3 Ostrich egg windowing and sham injection

The mean, standard deviation, minimum, and maximum for egg weight (g) of 89 Ostrich eggs were μ = 1466.2,

SD = 176.75, 432.1, and 1610.2, respectively (Figure 5). It should be noted that the maximum weight does not include measurements of 10 eggs that were rounded down to 1600 g when recorded. Of 36 windowed Ostrich eggs, one of the sham injected eggs (Group 2, i.e. air cell membrane kept intact; initial egg weight of 1412.7 g) hatched resulting in a healthy chick (Figure 6) that grew at a normal rate into adulthood. Another embryo survived up to the final stage of development but did not successfully pip, i.e. experienced mortality (Group 2 and with eggshell cap glued back on; initial egg weight of 1575.5 g). All other embryos experienced mortality earlier in development, i.e. on or before stage 32, based on chick embryo stages of development by Hamburger and Hamilton (1951).

Figure 4 Emu embryo development in a windowed egg (experienced mortality on day 25 of development, approximately 48–52 days required to hatch).

Figure 5 (a) Box and whisker plot of Ostrich egg (n = 89) weights for individual ostriches (n = 4), (b) box and whisker plot of the overall egg weights, and (c) histogram of overall egg weights (note: the maximum weight bin of 1600 g includes 10 eggs that were rounded down to 1600 g when recorded).


4. DISCUSSION

Based on the results, the null hypothesis that egg windowing with the subsequent hatching of chicks would not be possible for ratites, as in domestic Chickens and Turkeys, was rejected. Importantly, as far as we know, this is the first documentation of the successful hatching of a chick from a windowed ratite egg. Even though the efficiency was low, this result establishes that manipulating ratite embryos to create germline chimeras for genetic rescue or de-extinction is possible.

The group of Ostrich eggs with intact air cell membranes (Group 2) performed better compared to those without the air cell membrane left intact (Group 1). However, future research will be required to determine which factors may have been most responsible for successful development. Based on the results obtained herein, efforts to increase the survival of ratite chicks incubated in windowed eggs, and therefore reduce the number of embryos needed, will be necessary to increase the efficiency of avian transgenic and cloning projects given the financial costs involved.

In the present study, the efficiency of the Chicken windowing and embryo transfer to produce chicks using this method was lower compared to other studies. For example, Borwornpinyo et al. (2005) reported a higher percentage of survival and hatching, specifically, 75% ± 10.2% (windowing with Handi-Wrapâ) and 45.2% ± 13.8% (windowing with Saran Wrap), although embryos were placed into the recipient eggshell after the first three days of incubation and not two days after incubation, as in the present study. In hindsight, the early transfer of embryos to recipient eggshells with an artificial air space may have been detrimental to them by interfering with the physiological requirements concerning oxygen levels that change over time. Researchers at Crystal Bioscience (California, USA) have reported higher hatching success rates of windowed eggs with embryo transfers to surrogate eggshells (having artificial air spaces) after three days, thereby compensating for changing oxygen level requirements of the embryo (van de Lavoir and Mather-Love, 2006). The methodology in van de Lavoir and Mather-Love (2006) also includes adding 1 mL of tissue culture grade penicillin/streptomycin after the embryo transfer to thwart bacterial contamination. Incorporating

some of the methods being conducted at Crystal Bioscience might also help to improve future ratite egg windowing and embryo transfer results.

There are many other additional factors that may be responsible for the lower windowed Chicken egg viabilities in the present study compared to previous studies. Admittedly, a lack of previous experience with egg windowing was likely a contributing factor. There were also differences in the type of egg windowing materials used in some of the previous studies. Conspicuously, some of the windowed Chicken embryos survived up until the mid to later stages of development suggesting that the windowing or transfers themselves were not lethal, but likely due to some subsequent issue. The Humidaire incubator did not have a viewing window or internal video camera for external viewing of the eggs, as a result the incubator was opened and egg trays pulled out which was not ideal. It is possible that the exposure times from the daily opening of the incubator for the assessment of embryo development may have negatively influenced survival, as the room temperature was not the same as the incubator temperature.

An important factor in egg incubation and embryo development is egg weight loss. Determining the average weight loss for eggs with developing embryos over the incubation period can be useful for improving subsequent incubation outcomes for a particular species. Olsen and Olsen (1987) reported that weight loss for freshly laid eggs up until pipping is linear if the temperature and humidity are kept constant. Egg weight loss during incubation occurs when water vapour escapes from the egg by diffusing across the eggshell (Rahn and Ar,1974; Ar and Rahn, 1980), which depends upon the number of pores, pore structure and shell thickness (Ar et al., 1974; Rahn et al., 1976). If an egg begins losing weight faster than it should, or not fast enough, the humidity level of an incubator can be adjusted to slow or increase, respectively, egg water vapour conductance. However, in the present study adjustments for egg weight loss were not made and in future egg windowing experiments such adjustments might be implemented to improve hatching rates.

The importance of windowing eggs for access to the avian embryo goes beyond embryo transfer, e.g. transfer from a damaged eggshell to a non-damaged one, as it may

Figure 6 (a) Ostrich windowed eggs with eggshell cap glued back on after windowing, (b) windowed Ostrich egg with late stage of development embryo mortality (Group 2, sham injected with the inner air cell membrane left intact and with eggshell cap glued back on), and (c) Ostrich chick hatched from windowed egg (Group 2, sham injected with the inner air cell membrane left intact and without eggshell cap glued back on).


also allow for genetic rescue and de-extinction efforts intending to use germline chimeras. Notably, embryo transfer is not absolutely necessary to generate germline chimeras, which are generated by windowing the egg and injecting donor PGCs or genome editing transfection agents into the blood stream of the avian embryo during the developmental stage at which the embryo’s endogenous PGCs are circulating in the bloodstream prior to colonising the gonads. The production of intraspecific and interspecific germline chimeras can serve many purposes for avian conservation, for example, germline chimeras producing spermatozoa and ova from multiple PGC donors can reduce the number of breeding pairs necessary to conserve genetic diversity within a captive flock. Interspecific germline chimeras made using domestic species as PGC recipients could be used to produce offspring of PGC donor species that are difficult to propagate in captivity. The interspecific germline chimeras of highly fecund domestic PGC recipients could increase hatchling yields for population recovery of rare birds with low clutch sizes (van de Lavoir et al., 2012).

While embryo transfer may generally be unnecessary to produce germline chimeras, it is likely to be necessary when the expected offspring of germline chimeras are of a much larger size than the normal offspring of the surrogate germ-line chimera species (whether the difference in offspring/chimeric parent size is from genomic manipulation or the PGC donor species is a larger bird than the chimeric parent) – which will be the eventual case for de-extinction projects trying to generate proxies of extremely large extinct birds (i.e. Dodo, Great Auk, and Moa). It is unknown if an embryo genomically manipulated for gigantism will assume the egg-bound growth rate of the surrogate parent species and subsequently reach a larger size post-hatch or if the developing embryo will outgrow its egg space prior to hatch; in which case the amount of yolk reserve provided by the chimeric parent may also be an issue to be solved. Perhaps one way to solve the aforementioned issue would be with embryo transplantation methods in which the developing embryo is excised from its own yolk and grafted to a new yolk – a common practice for ex ovo embryo culture – that could be potentially modified to hatch extremely large chicks between interspecific chimeras, egg-shell surrogates, or eggshell and yolk surrogates (several newer methods of ex ovo embryo transplantation are outlined by Nagai et al. [2014]). No matter the modifications that may be required for successful de-extinction of giant birds, eggshell windowing and transfer will be universal steps for every avian de-extinction project, and as such further research should be conducted for any de-extinction candidate’s genomic template and potential chimeric parent and eggshell surrogate species. For ongoing de-extinction projects and future projects seriously being considered, similar embryo transfer experiments to those outlined in this paper could be conducted within genome-template and potential reproductive surrogate species for the Passenger

Pigeon (genomic template species = Band-tailed Pigeon, Patagioenas fasciata, possible chimeric parent = Rock Pigeon, Columba livia); Heath Hen (genomic template = Greater Prairie Chicken, Tympanuchus cupido, chimeric parent = domestic Chicken); Dodo (genomic template = Nicobar Pigeon, Caloenas nicobarica, chimeric parent or eggshell surrogate could be the domestic Goose, Anser anser domesticus), and Moa, (genomic template and chimeric parent candidate species = various Tinamou species, Emu, and Ostrich).

It should be mentioned that there currently is, and will continue to be, much debate regarding de-extinction and transgenic modifications of not only birds, but other species as well. Much consideration should be given before these projects are implemented and dialogue has already begun, and must continue, to take place among scientists, non-governmental organisations, and public and private entities. One has to consider not only the ability to de-extinct or create some proxy avian species, but also the behavioural, ecological, and ethical considerations of doing so.

Successful and efficient embryo transfer to new eggshells is the only means to pursue avian cloning unless reconstructed embryos are surgically implanted into surrogate mothers, which is deemed an invasive procedure and unjustifiable for endangered species unless an interspecific surrogate mother can be used. Enucleation of the avian ovum has been performed (Kjelland et al., 2014) but more work needs to be done for determining an appropriate electrofusion method, i.e. activation of reconstructed embryos, for a SCNT protocol and either a culture system involving eggshell culture or surgical embryo transfer techniques for the laying of cloned eggs. As stated in the introduction, successful avian SCNT protocols could offer advantages over germline transmission in some cases, especially if interspecies-SCNT could be achieved in birds as it has been for some wild mammalian species using domestic surrogate mothers (Sansinena et al., 2005; Gómez et al., 2009; Hwang et al., 2013).

5. ACKNOWLEDGMENTS

The authors would like to thank Ann Hoang, Jay Massey, and members of the Reproductive Sciences Laboratory; Dale Hyatt, Melvin Carter and staff at the Poultry Science Research, Teaching, and Extension Center; Lisa Roberts-Helton at the Winnie Carter Wildlife Center; Dr. George Stoica of the Department of Veterinary Pathobiology, College of Veterinary Medicine & Biomedical Sciences; Dr. Robert Burghardt of the Veterinary Integrative Biosciences Department, College of Veterinary Medicine & Biomedical Sciences; and Dr. Ian Tizard for their assistance with this project. This research was made possible by a grant, “Avian cloning to assist with preservation of endangered species: A pilot study” from the Schubot Exotic Bird Health Center


at Texas A&M University. All procedures performed in the present research were conducted in accordance with the Animal Care Guidelines of the Institutional Animal Care and Use Committee of Texas A&M University.

6. ELECTRONIC SUPPLEMENTARY INFORMATION

The ESI, Rhode Island Red Chicken embryo in windowed recipient Turkey eggshell during incubation (Video 1) and Rhode Island Red Chicken embryo pipping in windowed recipient Turkey eggshell with sterile petri dish cover (Video 2), is available through http://stl.publisher.ingentaconnect.com/content/stl/abr/supp-data/content-abr1600746_esi1 and http://stl.publisher.ingentaconnect.com/content/stl/abr/supp-data/content-abr1600746_esi2.

Published online: 7 July 2017

7. REFERENCES

Amadon, D. (1947) An estimated weight of the largest known bird. Condor, 49, 159–164.

Ar, A., Paganelli, C.V., Reeves, R.B., Greene, D.G. and Rahn, H. (1974) The avian egg: water vapor conductance, shell thickness, and functional pore area. Condor, 76, 153–158.

Ar, A. and Rahn, H. (1980) Water in the avian egg: overall budget of incubation. Amer. Zool., 20, 373–384.

Barrangou, R. (2014) RNA events. Cas9 targeting and the CRISPR revolution. Science, 344, 707–708.

Bengtson, S.-A. (1984) Breeding ecology and extinction of the Great Auk (Pinguinus impennis): Anecdotal evidence and conjectures. Auk, 101, 1–12.

Birkhead, T.R. and Nettleship, D.N. (1984) Egg size, composition and offspring quality in some Alcidae (Aves : Charadriiformes). J. Zool., 202, 177–194.

Borwornpinyo, S., Brake, J., Mozdziak, P.E. and Petite, J.N. (2005) Culture of chicken embryos in surrogate eggshells. Poult. Sci., 84, 1477–1482.

Brown, D. (1995) A guide to pigeons, doves, and quail, 1st edition. Australian Birdkeeper, South Tweed Heads, NSW Australia.

Cartwright, A.L. (2000) Incubating and hatching eggs. Document B-6092 of the Texas Agricultural Extension Service, Texas A&M University System, College Station, TX.

Dimitrov, L., Pedersen, D., Ching, K.H., Yi, H., Collarini, E.J., Izquierdo, S., van de Lavoir, M.-C. and Leighton, P.A. (2016) Germline gene editing in chickens by efficient CRISPR-mediated homologous recombination in primordial germ cells. PLoS ONE, 11, e0154303.

Gómez, M.C., Pope, C.E., Ricks, D.M., Lyons, J., Dumas, C. and Dresser, B.L. (2009) Cloning endangered felids using heterospecific donor oocytes and interspecies embryo transfer. Reprod. Fertil. Dev., 21, 76–82.

Hamburger, V. and Hamilton, H.L. (1951) A series of normal stages in the development of the chick embryo. J. Morphol., 88, 49–92.

Hipfner, J.M. (2000) The effect of egg size on post-hatching development in the Razorbill: an experimental study. J. Avian Biol., 31, 112–118.

Hwang, I., Jeong, Y.W., Kim, J.J., Lee, H.J., Kang, M., Park, K.B., Park, J.H., Kim, Y.W., Kim, W.T., Shin, T., Hyun, S.H., Jeung, E.B. and Hwang, W.S. (2013) Successful cloning of coyotes through interspecies somatic cell nuclear transfer using domestic dog oocytes. Reprod. Fertil. Dev., 25, 1142–1148.

Jefferey, J.S. (2001a) Emu production. Texas Agricultural Extension Service, The Texas A&M University System.

Jefferey, J.S. (2001b) Ostrich production. Texas Agricultural Extension Service, The Texas A&M University System.

Jeffrey, J.S., Martin, G.P. and Fanguy, R.C. (1996) The incubation of ratite eggs. Document L-5166 of the Texas Agricultural Extension Service, Texas A&M University System, College Station, TX.

Johnson, J.A., Altwegg, R., Evans, D.M., Ewen, J.G., Gordon, I.J., Pettorelli, N. and Young, J.K. (2016) Is there a future for genome-editing technologies in conservation? Anim. Conserv., 19, 97–101.

Johnson, J., Novak, B., Athrey, G., Shapiro, B., Phelan, R. and Brand, S. (2015) Whole genome sequence analysis reveals evolutionary history of extinct Heath Hen. In: Joint Meeting of the American Ornithologist’s Union & Cooper Ornithological Society, 1 August, Norman, Oklahoma.

Kjelland, M.E., Blue-McLendon, A. and Kraemer, D. (2012) Determining air cell location and embryo development in opaque shelled eggs. Avian Biol. Res., 5, 99–102.

Kjelland, M.E., Romo, S. and Kraemer, D. (2014) Avian cloning: adaptation of a technique for enucleation of the avian ovum. Avian Biol. Res., 7, 131–138.

Lai, Y.Y. and Liu, J. (2015) Transparent soft PDMS eggshell. Sci. China Technol. Sc., 58, 273–283.

Livezey, B.C. (1993) An ecomorphological review of the Dodo (Raphus cucullatus) and Solitaire (Pezophaps solitaria), flightless columbiformes of the Mascarene Islands. J. Zool., 230, 247–292.

McGrew, M.J., Sherman, A., Ellard, F.M., Lillico, S.G., Gilhooley, H.J., Kingsman, A.J., Mitrophanous, K.A. and Sang, H. (2004) Efficient production of germline transgenic chickens using lentiviral vectors. EMBO Rep., 5, 728–733.

Mitchell, K.J., Llamas, B., Soubrier, J., Rawlence, N.J., Worthy, T.H., Wood, J., Lee, M.S.Y. and Cooper, A. (2014) Ancient DNA reveals elephant birds and kiwi are sister taxa and clarifies ratite bird evolution. Science, 344, 898–900.

Moum, T., Arnason, U. and Arnason, E. (2002) Mitochondrial DNA sequence evolution and phylogeny of the Atlantic Alcidae, including the extinct great auk (Pinguinus impennis). Mol. Biol. Evol., 19, 1434–1439.

Nagai, H., Mak, S.S., Weng, W., Nakaya, Y., Ladher, R. and Sheng, G. (2011) Embryonic development of the emu, Dromaius novaehollandiae. Dev. Dyn., 240, 162–175.

Nagai, H., Sezaki, M., Nakamura, H. and Sheng, G. (2014) Extending the limits of avian embryo culture with the modified Cornish pasty and whole-embryo transplantation methods. Methods, 66, 441–446.

Nicholls, H. (2012) The legacy of Lonesome George: tortoise’s death spurs Galapagos conservation efforts. Nature, 487, 279–280.


Olsen, P.D. and Olsen, J. (1987) Egg weight loss during incubation in captive Australian Kestrels Falco cenhroides and Brown Goshawks Accipiter fasciatus. Emu, 87, 196–199.

Pennisi, E. (2013) The CRISPR craze. Science, 341, 833–836.Perry, M.M. (1988) A complete culture system for the chick embryo.

Nature, 331, 70–72.Preston, F.W. (1968) The shapes of bird’s eggs: mathematical

aspects. Auk, 85, 454–463.Rahn, H. and Ar, A. (1974) The avian egg: incubation time and

water loss. Condor, 76, 147–152.Rahn, H., Paganelli, C.V., Nisbet, I.C.T. and Whittow, G.C. (1976)

Regulation of incubation water loss in eggs of seven species of terns. Physiol. Zool., 49, 245–259.

Rowlett, K. and Simkiss, K. (1987) Explanted embryo culture: in vitro and in ovo techniques for domestic fowl. Br. Poult. Sci., 28, 91–101.

Sang, H. and Sherman, A. (2003) Method of preparing eggs for nuclear transfer and uses thereof WO2003049537 (application number). United States.

Sang, H. and Sherman, A. (2005a) Method of preparing eggs for nuclear transfer and uses thereof. US20050015825. United States.

Sang, H. and Sherman, A. (2005b) Method of preparing eggs for nuclear transfer and uses thereof. EP1455573. Europe.

Sang, H. and Sherman, A. (2005c) Method of preparing eggs for nuclear transfer and uses thereof. 2002356292 (application number). Australia.

Sansinena, M.J., Hylan, D., Hebert, K., Denniston, R.S. and Godke, R.A. (2005) Banteng (Bos javanicus) embryos and pregnancies produced by interspecies nuclear transfer. Theriogenology, 63, 1081–1091.

Shapiro, B. (2016a) Pathways to de-extinction: how close can we get to resurrection of an extinct species? Funct. Ecol., https://doi.org/10.1111/1365-2435.12705.

Shapiro, B. (2016b) The curious case of the Dodo: Leveraging the Nicobar pigeon genome to resurrect this long-extinct bird. In: Plant and Animal Genomes Conference XXIV, 11 January, San Diego, CA.

Shapiro, B., Green, R., Soares, A.E.R., Novak, B. and Corbett-Detig,

R. (2016) Population genomics of Passenger Pigeons. In: Plant

and Animal Genomes Conference XXIV, 10 January, San Diego,

CA.

Shapiro, B., Sibthorpe, D., Rambaut, A., Austin, J., Wragg, G.M.,

Bininda-Emonds, O.R.P., Lee, P.L.M. and Cooper, A. (2002)

Flight of the Dodo. Science, 295, 1683.

Tyack, S.G., Jenkins, K.A., O’Neil, T.E., Wise, T.G., Morris, K.R.,

Bruce, M.P., McLeod, S., Wade, A.J., McKay, J., Moore, R.J.,

Schat, K.A., Lowenthal, J.W. and Doran, T.J. (2013) A new

method for producing transgenic birds via direct in vivo

transfection of primordial germ cells. Transgenic Res., 22,

1257–1264.

van de Lavoir, M.-C., Collarini, E.J., Leighton, P.A., Fesler, J., Lu,

D.R., Harriman, W.D., Thiyagasundaram, T.S. and Etches, R.J.

(2012) Interspecific germline transmission of cultured primordial

germ cells. PLoS ONE, 7, e35664.

van de Lavoir, M.-C., Diamond, J.H., Leighton, P.A., Mather-Love,

C., Heyer, B.S., Bradshaw, R., Kerchner, A., Hooi, L.T., Gessaro,

T.M., Swanberg, S.E., Delany, M.E. and Etches, R.J. (2006)

Germline transmission of genetically modified primordial germ

cells. Nature, 441, 766–769.

van de Lavoir, M.-C. and Mather-Love, C. (2006) Avian embryonic

stem cells. Methods Enzymol., 418, 38–64.

Véron, N., Qu, Z., Kipen, P.A.S., Hirst, C.E. and Marcelle, C. (2015)

CRISPR mediated somatic cell genome engineering in the

chicken. Dev. Biol., 407, 68–74.

Zuo, Q., Wang, Y., Cheng, S., Lian, C., Tang, B., Wang, F., Lu, Z.,

Ji, Y., Zhao, R., Zhang, W., Jin, K., Song, J., Zhang, Y. and Li, B.

(2016) Site-directed genome knockout in chicken cell line and

embryos can use CRISPR/Cas gene editing technology. G3, 6,

1787–1792.

manipulating the avian egg: applications for embryo transfer, … · 2017-10-01 · avian biology...

Documents