Artificial insemination technology for ratites: a review

Review CSIRO PUBLISHING Australian Journal of Experimental Agriculture, 2008, 48, 1284–1292 www.publish.csiro.au/journals/ajea Artificial insemination technology for ratites: a review I. A. Malecki A,C, P. K. Rybnik B and G. B. Martin A AInstitute of Agriculture M092, Faculty of Natural and Agricultural Sciences, University of Western Australia, Crawley, WA 6009, Australia. BDepartment of Animal Sciences, Institute of Genetics and Animal Breeding, Polish Academy of Sciences, Jastrzebiec, 05-552 Wolka Kosowska, Poland. CCorresponding author. Email: imalecki@animals.uwa.edu.au Abstract. In ratite farming, the low male to female ratio in the mating system restricts genetic improvement and prevents reduction of the number of males kept on-farm for fertilisation of the female flock. These issues can be overcome and the industry can better realise its potential by using artificial insemination (AI) technology. It is the only practical method for intensive genetic improvement of reproduction and the production of eggs, chicks, oil, meat and leather. For AI to be feasible, we need reliable methods for semen collection, artificial insemination, prolonged storage of spermatozoa in the female tract, high rates of lay, efficient protocols for semen storage, and a panel of quantitative methods for measuring true fertility and hatchability, sperm supply rates in vivo and sperm viability in vitro. For both emus and ostriches, prolonged sperm storage in females has already been demonstrated. Methods for semen collection and artificial insemination, using animal-friendly techniques, have also been developed. Semen storage and cryopreservation protocols are yet to be optimised and we still need to overcome the male-dependent rate of lay, but adoption of AI technology by the ratite industries is now feasible. It also seems likely that these technologies will be relevant to wild ratites that need intensive conservation efforts, such as cassowaries, rheas and ostrich subspecies. Additional keywords: birds, breeding, Dromaius, Struthio, sexual behaviour. Introduction Ratite species such as the ostrich (Struthio camelus), emu (Dromaius novaehollandiae) and rhea (Rhea americana) are fundamentally attractive for farming for the production of leather, meat, oil and feathers, yet ratite farming has been established with variable success, due to adverse climatic conditions, poor management, insufficient infrastructure and poor genetics. Challenges posed by the biology of the species and inadequate management practices have led to problems such as variable egg production, fertility, hatchability, embryo mortality, and low chick survival and poor growth rate. Many farms attempt to solve problems in production efficiency by changing their management practices or by applying the latest research outcomes, but progress is very slow and often geographically isolated, rather than on national or global industry scales. Without doubt, there have been advances in almost every area of production, but progress is too slow for the industry to be commercially attractive, let alone expanding; even the largest and most advanced of them, the ostrich industry, is still inefficient (Van Rooyen et al. 1998). Major improvements will depend on a better understanding of ratite biology, as well as opportunities offered by new technologies in breeding and genetics that will advance the industry beyond traditional farming (Cloete et al. 1998; Sales 2006, 2007). Worldwide, the ratite industries lack structured breeding programs that would guarantee rapid genetic improvement. This is in sharp contrast with other livestock industries, especially poultry, pigs and dairy cattle, where striking increases in © CSIRO 2008 productivity have been achieved in a relatively short period of time. Moreover, the ratite industries still rely on natural reproduction, leading to a major biological constraint – the dominant influence of male–female pair-bonding in the mating system. The outcome is the need for farmers to maintain a low male:female ratio. Apart from the cost of maintaining a large number of males, natural mating hampers rapid genetic improvement because the genes of an elite male can be offered to only a few females. In colonies, the males may mate with a greater number of females, but there is no guarantee that males with the highest genetic merit would pass on their genes and certainly nowhere near the selection pressure that is normally applied in other animal industries. To achieve rapid and sustained genetic improvement, the ratite industries need to adopt advanced reproductive technologies in: semen collection, fertility assessment, semen handling, storage and cryopreservation, and artificial insemination (AI). This review focuses on recent developments in this area for emus and ostriches, and supports the view that it is very feasible for these industries to adopt these technologies now. Semen collection In an AI program, semen is required regularly so collection methods need to guarantee high quality ejaculates with an optimum frequency from elite males. This requires a welldefined protocol for semen collection that is both animal- and human-friendly and will yield quality ejaculates. Good 10.1071/EA08141 0816-1089/08/101284 Artificial insemination technology for ratites equipment is clearly necessary but, in addition, we need a strong knowledge of animal behaviour, including the ability to predict adult sexual behaviour during the prepubertal phase of bird development. Semen collection methods Until recently, the only method for collecting semen from ostriches was manual massage (Von Rautenfeld 1977; Bertschinger et al. 1992; Irons et al. 1996; Hemberger et al. 2001), but this was labour intensive and unreliable, so it has not been adopted by the industry. In this method, the male has to be confined in a ‘V’ shaped crush to prevent injury to handlers and the bird. Once restrained and settled, the male copulatory organ, the phallus (King 1981b), is extruded out of the cloaca and held down using a cloth for allowing firm grip. The fingers of the other hand apply a gentle massage of the semen papillae area until semen flows down the phallic groove. There are now reliable methods for inducing ejaculation into an artificial cloaca (AC) for both ostriches and emus. Males can be trained to one of two methods, either using a ‘teaser’ female or using a ‘nonteaser’ approach for males that have been imprinted to direct their courtship towards humans. The teaser method follows the normal sequence of courtship and mating behaviour – the male follows the female in a courtship display and, upon submission, the male mounts the female (Malecki et al. 1997a; Ya-jie et al. 2001; Rozenboim et al. 1999, 2003; Rybnik et al. 2007). The phallus is redirected into an AC into which the male is induced to ejaculate. Some of the critical factors that determine the success of the teaser method in the ostrich have been described (Rozenboim et al. 2003; Rybnik et al. 2007), but the method is not yet optimal because the daily ejaculate output cannot be controlled and the optimum collection frequency is not yet known. In the emu, this information is available and the best output is achieved by separation of sexes after semen collection (Malecki et al. 1997b) so the male has no opportunity to ejaculate until next collection. In the nonteaser method, males are trained by taking advantage of courtship behaviour directed towards humans. A human directed courtship has been described by Malecki et al. (1997a, 1997b) in emus and by Bubier et al. (1998) and Rozenboim et al. (2003) in ostriches. In training, when desirable human–bird interaction stimulates the males to express their normal sexual behaviour, their courtship is returned by giving them an opportunity to mount the arm and shoulder of the semen collector, in the case of the emu (Malecki et al. 1997a), or a dummy, in the case of the ostrich (Malecki and Martin 2005b; Rybnik et al. 2007). The male emu squats in front of the semen collector and, while moving on his hocks forward, attempts mounting. The collector responds by squatting next to the male and placing the AC on the phallus to induce ejaculation. In the ostrich, the male squats and performs kantling display, then stands up, mounts the dummy placed between him and the collector, and ejaculates into the AC mounted inside the dummy. Selection of teaser females In the emu and ostrich, teaser females play a critical role in collection success, ejaculate repeatability, and maintenance of Australian Journal of Experimental Agriculture 1285 male libido, so it is very important to select a good teaser. Studies of this are, however, limited to a few reports on selection criteria and the apparent characteristics of good teaser females (Malecki et al. 1997a; Rozenboim et al. 1999, 2003; Rybnik et al. 2007), but there is lack of quantitative data. The initial selection is based on readiness to sit for a human and the subsequent display of normal sexual behaviour so that the teaser female elicits good responses from a given male. This is particularly important for shy temperament males. The crouching behaviour is essential because this is a potent stimulus for the male to mount the female. Subsequently, the teaser needs to remain in a crouching position until ejaculation is complete. Selection of the teaser is complete after testing the female’s response to a human and then to a male. In the emu, the size and temperament of the females requires particular consideration because, in this species, the female is generally larger than a male and she appears to play a dominant role in the natural mating system (Blache et al. 2000). Some teasers can show preferences for particular males – this is unavoidable, so several teaser females are needed and used interchangeably when required. It is yet to be demonstrated that separation of the sexes, as used in the emu teaser method, is useful for the ostrich. For ostriches, additional female criteria may need to be identified. Semen collection equipment The equipment for collecting semen depends on the method of semen collection but, in general, it comprises the AC and the dummy female (Malecki et al. 1997a; Rybnik et al. 2007). In the teaser method, only the AC is used. For the emu, the AC is made of a plastic tube ~16 cm long and 5 cm in diameter with an internal rubber liner (as used for rams). The space between the tube and the liner is filled with warm water at ~40°C (cloacal temperature is ~38°C). A plastic funnel with a collecting vial is mounted at the end of the tube. For the ostrich, the AC can be made of a PVC tube 13 cm in diameter and 35 cm long, and the rubber liner can be that used for horses (A/V Colorado Liner; Pacific Vet Pty Ltd, Vic.). The space between the tube and a liner is again filled with warm water at 40°C. A collecting vial is attached to a rubber cone mounted on one end of the AC. For the AC to be used in the dummy method, three strips of firm dense foam can be glued on the outside surface of the PVC tube lengthwise to allow a tight fit into the PVC tube inside the dummy. A dummy female is not required for the emu when using a nonteaser approach because, when mounting a female, the male emu keeps both legs on the ground. For this method the human collector plays the role of dummy because the male is given access to the collector’s arm and back for mounting, while the collector kneels on the ground (Malecki et al. 1997a). In the ostrich, the male mounts the female by putting his leg on her back, so a dummy is needed. In our initial studies (Malecki and Martin 2005b), the dummy was made from a plastic drum cut in half and a wooden structure covered with a canvas was added for leg support. Subsequently, Rybnik et al. (2007) used a hemp sack filled densely with wheat straw and stitched at the end. Inside, there was a wooden structure that supported a built-in PVC tube of 15 cm in diameter and provided firm leg support for the male. The tube is fitted 3 cm above the ground level and the AC is inserted into that tube. 1286 Australian Journal of Experimental Agriculture I. A. Malecki et al. Frequency of collection and semen output Emu and ostrich ejaculates collected by the AC show low variability in mass motility and are within the normal range for fresh avian semen. Both methods, teaser and nonteaser, yield quality ejaculates (Table 1). Production of semen and spermatozoa can be affected by collection frequency, but the frequency of collection can depend on the method of collection itself. Naturally, male birds can mate more than once a day and, with a method that elicits natural male responses, frequency of semen collection can maximise the output without adverse effects on the male. Semen collection by manual massage is generally less frequent than once a day (Hemberger et al. 2001). Daily collections can be carried out in emus and ostriches. If collection frequency is increased to twice daily in the emu, the output of semen and spermatozoa is doubled over a 6-day period, but longer durations have not been investigated (Malecki et al. 1997b). Ejaculates have been collected daily from emus for 16 days (Malecki et al. 1997b) and from ostriches for 10 days (Rybnik et al. 2008b) and no adverse effect on male behaviour or semen output was observed. The effect of frequencies higher than once a day has not yet been studied in the ostrich. The proportions of live and normal spermatozoa tend to be lower for the manual massage (Bertschinger et al. 1992; Hemberger et al. 2001) than for the teaser (Ya-jie et al. 2001; Rozenboim et al. 2003; P. K. Rybnik, unpubl. data) or nonteaser methods (P. K. Rybnik, unpubl. data). Age would be expected to affect the production of semen and spermatozoa in ratites because males can take up to 3 years to mature. Thus, 2-year-old emus had lower ejaculate volume and sperm concentration than 3–5-year-olds (Malecki 1997). In the ostrich, semen parameters were similar in 2- and 3-year-old males (Rybnik et al. 2008a) and were comparable to those collected from 7–8-year-olds, although our data are limited to two individuals. In emus and ostriches, high between-male variation has been observed in parameters for collected ejaculates and the likely confounding factors are genotype, temperament, paddock position, year, month, timing of puberty, libido, method of collection and the handler or semen collector. It is yet to be confirmed, but we expect that, in the ostrich, collection at frequencies of more than once a day would yield more semen than single daily collections, although this might depend on the method of collection and male libido, factors that Table 1. have already been identified in the emu (Malecki et al. 1997b) and observed in the ostrich (I. A. Malecki and P. K. Rybnik, unpubl. data). Determining the optimal collection frequency is essential for developing the most efficient use of semen, maximising semen extension, and reducing the number of males in an AI system. Fertility technology Until recently, emu and ostrich breeding lacked objective, quantitative methods for determining fertility. Fertility has a direct effect on reproductive efficiency and a range of approaches are needed for evaluation of fertility, whether emus and ostriches are under natural mating or AI, in order to detect birds with fertility problems and to identify elite breeding birds or stock for fertility traits (Malecki and Martin 2002b, 2003). Measuring fertility Development of approaches for evaluation of emu and ostrich fertility has been based on the same principles used for poultry (Wishart 1995; Wishart and Staines 1995). A range of techniques can be used from egg break-out to perivitelline techniques and sperm–egg interaction assay in vitro. Egg break-out fertility Candling of eggs is unreliable because clear eggs, where no embryonic development is detected due to very early embryonic death, can be misdiagnosed and true fertility underestimated. Thus, the egg needs to be opened and the status of the germinal disc (GD) determined. After the egg is open, the GD is located with the unaided eye, usually on top of the floating egg yolk. The egg white overlaying the yolk is removed to expose the intact yolk and the GD region. With experience, observations can be made with the unaided eye but, for detailed observation, a dissecting microscope is essential. The GD is classified as either unfertilised (blastodisc) or fertilised, when it contains the blastoderm. The blastodisc consists of a group of white yolk droplets surrounded by a yellow ring, within which numerous vacuoles can be found. The fertilised GD (blastoderm) generally consists of two areas, a clear concentric one in the centre, the area pellucida, surrounded by an opaque ring, the area opaca (Romanoff and Romanoff 1949; Gupta and Bakst 1993; Bakst et al. 1998). The area pellucida is usually clearly delineated from the area opaca and contains no vacuoles or white yolk Characteristics of normal emu and ostrich ejaculates collected by teaser and nonteaser methods Species Volume (mL) Emu Ostrich Ostrich Ostrich 0.2–1.6 0.1–1.1 0.2–2.0 0.5–2.9 2.2–6.3 2.1–8.2 1.6–4.8 2.0–6.8 Teaser method 79–95 93–96 87–93 82–98A I. A. Malecki (unpubl. data) Rozenboim et al. (2003) Ya-jie et al. (2001) Rybnik et al. (2007, 2008a, 2008b) Emu Ostrich 0.1–1.0 0.2–2.0 1.8–6.5 1.5–6.7 Nonteaser method 75–96 78–96A I. A. Malecki (unpubl. data) Rybnik et al. (2007, 2008a, 2008b) AP. Conc. of sperm Live sperm (×109/mL) (%) K. Rybnik (unpubl. data). Reference Artificial insemination technology for ratites droplets. Within the area pellucida, clusters of small white cells may be visible, depending on the stage of blastoderm development (Malecki et al. 2005d). Perivitelline techniques Interactions between sperm and egg in vivo and in vitro can provide very important evidence about the fertility status of males and females, and can help with predicting fertility and the probability of fertilisation (Wishart 1997). Assessment is based on sperm populations in the perivitelline layer of the egg yolk membrane. One population leaves holes in the inner perivitelline layer of the vitelline membrane (IPVL) caused by successful hydrolysis, the other population has remained outside the ovum, trapped in the outer perivitelline layer of the vitelline membrane (OPVL). Counts of IPVL holes provide evidence about the success of penetration and fertilisation and, when these data are added to the number of OPVL sperm, information is provided about the success of mating, the ability of the female to transport sperm, the ability of the sperm to reach the egg, and the ability of the sperm to penetrate the membrane. The application of those in vivo interactions for evaluation of fertility in emus and ostriches has been presented in detail elsewhere (Malecki and Martin 2005b), so here we provide only a brief summary. Sperm–egg interaction assay in vitro An assessment of male and female fertility in vitro is carried out using a homologous sperm–egg interaction (S-E) assay as developed for the emu (Malecki et al. 2005b). A similar assay is yet to be developed for the ostrich. In this assay, the number of holes (points of hydrolysis) made by sperm in the IPVL is used to identify males with superior sperm fertilising ability and females whose eggs are ‘attractive’ to sperm. On the other hand, males with inferior fertilising ability and the females with poor sperm penetration of their egg membrane can be identified. It is also a useful assay for measuring sperm fertilising ability following short-term storage and cryopreservation. Sperm supply, storage, sperm loss and fertile period as parameters of fertility The efficiencies of sperm supply by males and sperm use by females are two of the most critical factors affecting fertilisation rates. Our understanding of sperm supply, transfer and storage in vivo, and the maintenance of high efficiency of sperm use under natural mating have improved, and this knowledge could be applied to the development of AI technology (Malecki and Martin 2005b). In birds, the number of sperm that the female can receive depends on copulation frequency and on ejaculate quality (Birkhead 1988; Cornwallis and Birkhead 2007); ratites are no exception. In the pair mating system, compatible emus and ostriches can show good fertility, but ostrich mating systems with more than one female can demonstrate variable fertility (Malecki and Martin 2005b). Differences in the mating systems of ostriches suggest that the males preferentially allocate sperm to females, or that females may control ejaculate utilisation. Variability in sperm supply to sperm storage sites could affect the female fertile period. Fertility technology can assist in identification of subfertile males or females. Highly fertile birds could be retained in Australian Journal of Experimental Agriculture 1287 naturally mated flocks or be selected for an AI program. There might be a commercial advantage from understanding mate choice and sperm allocation in both emus and ostriches. Female emus and ostriches store sperm in sperm storage tubules (SSTs) and release them over a period of days or weeks to fertilise a sequence of ova (Bezuidenhout et al. 1995; Malecki and Martin 2002a; Malecki et al. 2004). Emus and ostriches store sperm for a prolonged time, despite the constant presence of the male and their ability to mate for every egg. In avian species in general (Bakst et al. 1994), the duration of sperm storage is related to the sperm storage capacity of the female oviduct. Two sperm storage sites are important to the female: the sperm storage tubules of the utero–vaginal junction and the infundibular crypts (Bakst et al. 1994) – the crypts have not been found in the emu or ostrich. SSTs have been described in the emu and ostrich oviduct and, in contrast with the emu, the ostrich SSTs appear to form a more extensive network of tubules (Bezuidenhout et al. 1995; Holm et al. 2000; Malecki 1997). Also, the ostrich utero–vaginal junction is larger than that of the emu. Together, these factors suggest that female ostriches can store more sperm than female emus. Greater capacity for reception of sperm by female ostrich than the female emu suggests that the male ostriches can supply more sperm than male emus. This is supported by evidence from single ejaculates (Malecki et al. 1997b; Ya-jie et al. 2001; Rozenboim et al. 2003; Rybnik et al. 2007), although the large numbers of sperm that the female ostrich receives could be the result of a high frequency of sperm supply in this communally breeding bird. Further evidence was obtained from direct comparison of emu and ostrich eggs (Malecki and Martin 2003), showing more sperm in the perivitelline membrane above the GD in the ostrich than in the emu. The rates of sperm loss in the emu (27% per day) and ostrich (17% per day) are relatively slow when compared with that in poultry and a few other species, except for the turkey (Malecki and Martin 2002b). This is important for the ratites because consecutive ovulations occur 2 (ostrich) or 3 days (emu) apart, whereas sperm appears to be released constantly from avian SSTs (Bakst et al. 1994). Therefore, female emus and ostriches have the potential to produce a relatively good number of fertilised eggs within their fertile period (Malecki and Martin 2005a). Fertile period Determination of the fertile period is a critical prerequisite to determining the optimum frequency for AI. Female emus and ostriches have a similar mean duration of the fertile period of just over 2 weeks, during which we would expect the emu to produce up to six fertilised eggs and the ostrich to produce up to eight (Malecki and Martin 2002a, 2002b; Malecki et al. 2004). However, these are average values so the maintenance of maximum flock fertility after AI is likely to require an insemination interval of less than 2 weeks because of betweenfemale variation in the duration of the fertile period, which could mean less fertilised eggs. On the other hand, in future, the optimum frequency for AI may decrease through selection for improved duration of the fertile period (Brillard et al. 1998) and more fertilised eggs gained per AI. 1288 Australian Journal of Experimental Agriculture Artificial insemination Seasonality of breeding High yields of semen and spermatozoa throughout the period of egg production are essential for efficient production but, in both emus and ostriches, the production of sperm and eggs is affected by season. The emu is a short-day breeder and lays only over the autumn and winter months (Malecki et al. 1998b), whereas the ostrich is a spring–summer breeder, irrespective of the latitude (Degen et al. 1994; Malecki and Martin 2003; Gee et al. 2004). The breeding season lasts for 5–6 months in the emu and for 7–8 months in the ostrich, but there appears to be considerable between-bird variation in both males and females in the commencement and termination of gamete production (Malecki et al. 1998b; Malecki and Martin 2000, 2003; Rybnik et al. 2008b). AI technology could play a major role in removing some of that variation. The seasonal timing of reproduction is primarily controlled by photoperiod in the emu, with rain being a modulator (Williams et al. 1997; Blache et al. 2001), whereas factors such as socio-sexual signals, diet and photoperiod seem to play a significant role in the ostrich (Bunter et al. 2001; Lambrechts et al. 2005). Factors affecting the timing of reproduction have not yet been sufficiently studied or explored for the purpose of improving reproductive efficiency and this needs to be a high priority for future research. Inter-oviposition intervals and the ovulation cycle On average, ostriches lay one egg every 2 days, whereas emus lay one egg every 3 days (Malecki and Martin 2002a; Bronneberg et al. 2005). In a succession of ovipositions, the interval is variable and there are pauses that may last days, sometimes weeks. This pattern of lay shows that both species lay eggs in clutches. This means that there is scope for selection for long clutch duration and for pauses that are short or nonexistent as we aim to achieve maximum egg output. Progress in this area will also lead to more efficient use of sperm and to maximisation of the number of eggs per AI. In the ostrich, separation of the sexes quickly reduces the rate of lay (Cloete et al. 1998; Malecki et al. 2004). In the emu, this problem is smaller provided the male remains in visual proximity of the female (Malecki and Martin 2002b). However, in both species, these effects are not observed in all females so there is an opportunity to select for females that are less affected by separation from males, provided the basis for the phenomenon is understood. On the other hand, recent evidence suggests that rearing conditions may affect future mate preferences: prepubertal female ostriches were identified as friendly to humans and then further selected based on their crouching behaviour in response to humans and then maintained as a female-only flock. These females laid eggs and could be artificially inseminated following a voluntary crouch (Malecki et al. 2008). As suggested by Malecki et al. (1997a) and Bubier et al. (1998), sexual preference for humans in both emus and ostriches could result from imprinting on humans soon after hatching and, if the phenomenon is associated with laying, selective breeding could take advantage of the opportunity. Timing of AI It is generally accepted that inseminations performed near the time of oviposition result in a loss of too much sperm (Bakst I. A. Malecki et al. et al. 1994). In the emu, if AI is carried out in the morning of day 3 of the egg cycle (day of oviposition), the duration of fertility is reduced (Malecki and Martin 2004). This may also apply to ostriches, indicating that inseminations should not be performed near oviposition time. Given that laying starts early afternoon, female ostriches may need to be inseminated the next day, but the optimum AI time remains to be determined in future studies. AI dose The optimal dose of semen for AI has been investigated in the emu (Malecki and Martin 2004) but not the ostrich. In the emu, 200 million freshly collected sperm are needed for fertilisation of three eggs laid every 3 days. A dose for stored or frozenthawed sperm has not yet been determined. Female ostriches may require larger doses, but more eggs should be produced because of the shorter duration of the egg cycle. Factors such as semen handling and storage, timing of insemination, insemination technique and efficiency of sperm use by females, will affect numbers of sperm required for insemination and these topics await future research. AI techniques and equipment Artificial insemination has been carried out in emus and ostriches and chicks have been produced. The methods appear to be reliable, although there is still room for improvement. Female emus develop a responsive behaviour and crouch sexually, thus allowing insemination to be carried out without restraining the birds (Malecki and Martin 2004). Until recently, female ostriches needed to be caught and restrained for this procedure, which requires up to four people (Von Rautenfeld 1977). A behavioural approach, as developed for female emus, has also been used for female ostriches but responsive females had to be identified and accustomed to this procedure (Malecki and Rybnik 2008). In both species, semen can be deposited into the vagina despite anatomical differences in their cloacae (King 1981a). For inseminating the female emu, an insemination straw of medium rigidity and length 15 cm is mounted on a tuberculin syringe and, with the aid of a speculum, the lumen of the cloaca is widened so the straw can be inserted into the vagina. In the ostrich, the insemination straw needs to be more rigid than for the emu and ~30 cm long. It is inserted into the cloaca and guided by the fingers of one hand alongside the female phallus into the vaginal orifice and then further into the vagina. It appears that these techniques ensure most semen being retained in the oviduct for storage and fertilisation of subsequent ova. This seems to maximise efficient use of the semen dose but full evaluation of all procedures is still needed for the ostrich. Semen holding and freezing The handling and processing of semen affect its quality and therefore underpin the future success of AI in ratites. Semen collected by the teaser and nonteaser methods is of good quality and is suitable for storage (Malecki et al. 1997b, 1998a; Malecki et al. 2005a; Rybnik et al. 2007), but the loss of sperm quality during short-term storage and freezing (cryopreservation) can be considerable. Developing an optimal diluent is crucial to the maintenance of sperm quality during the period between semen collection and AI. Several diluents have already been investigated. For short- Artificial insemination technology for ratites term storage (hours to days), emu and ostrich semen have been tested with the synthetic diluents used for poultry semen. Emu semen has been diluted with Lake’s diluent, BPSE (Sexton and Fewlass 1978), NaCL-TES (Bootwalla and Miles 1992) and modified versions of Lake’s (Lake 1960) diluent, termed UWAEmu 1, 2 and 3 (Malecki and Martin 2000). Ya-jie et al. (2001) investigated several diluents for storage of ostrich spermatozoa and found MEM (Minimal Essential Medium) to be the most effective. Ostrich semen was also diluted with EK (Lukaszewicz 2002), Lake’s and 0.85% NaCl diluents (I. A. Malecki, P. K. Rybnik, A. Kowalczyk, E. Lukaszewicz and H. Naranowicz, unpubl. data). Short-term semen storage Ya-jie et al. (2001) found that ostrich spermatozoa could survive at 5°C for up to 72 h when diluted with MEM, but only up to 24 h in 0.85% NaCl, although only the maximum time of sperm survival was estimated so the daily rate of sperm loss is not known from this study. Changes in motility, expressed as proportions of progressively motile sperm (Allen and Champion 1955), and in proportions of live spermatozoa (the dye-exclusion test using nigrosine-eosine [N-E] staining; Lake and Stewart 1978) were investigated by storing diluted ostrich semen at 5°C in EK, Lake’s and NaCl diluents. Semen was diluted 1 part to 3 or 6 parts of diluent (v/v). The proportions of Table 2. Maximum maintenance (hours) of emu and ostrich sperm viability as estimated by various tests of sperm quality Medium DilutedB Undiluted DilutedC DilutedB Undiluted Time 1289 live spermatozoa in EK diluent did not change in 24 h and only 25% loss was detected after a further 24 h of storage (Table 2). Poorer survival was recorded in Lake’s and NaCl. Sperm survival was better if semen was diluted 1:6 than 1:3. Motility in EK diluent was maintained for 12 h and declined by 10% in the next 12 h. Decline over the next 24 h was more rapid. The loss of motile sperm was more pronounced in Lake’s and NaCl diluents (I. A. Malecki, P. K. Rybnik, A. Kowalczyk, E. Lukaszewicz and H. Naranowicz, unpubl. data). In the emu, sperm survival was first studied in BPSE and Lake’s diluents over 24 h at 5°C. The proportions of live spermatozoa (N-E test) and motility declined by 15% in 24 h (Malecki and Martin 2000). Subsequently, we developed tests based on hypo-osmotic swelling (HOS) and sperm-egg membrane hydrolysis (S-E) and showed that, for the emu, sperm membrane integrity and fertilising ability were maintained only for 6 h when stored diluted at 4°C (Malecki et al. 2005a, 2005b, 2005c). As estimated by the HOS and S-E tests, better survival of emu spermatozoa was achieved at 20°C than at 4°C. Sperm function was maintained for 24 h when semen was diluted with ‘emu’ diluents (Malecki and Martin 2000) or for 6–24 h if stored undiluted (Table 2). In poultry diluents, sperm function is only maintained for 6 h at 4°C, an outcome similar to that observed with storage in these diluents at 20°C (Malecki et al. 2005c). In the field, there have been no insemination trials yet with stored emu or ostrich spermatozoa, so we cannot relate laboratory results with egg fertility or duration of sperm storage in the female tract, but it appears that, for emu semen, storage at 20°C Sperm quality test Emu sperm stored at 4°CA 6 Swelling (HOS) 6 Membrane hydrolysis (S-E) 6 Dye exclusion (N-E) <6 Swelling (HOS) <6 Membrane hydrolysis (S-E) 6 Dye exclusion (N-E) Emu sperm stored at 20°C 24 Swelling (HOS) 24 Membrane hydrolysis (S-E) 48 Dye exclusion (N-E) 6 Swelling (HOS) 6 Membrane hydrolysis (S-E) 6–24 Swelling (HOS) 6 Membrane hydrolysis (S-E) 24 Dye exclusion (N-E) DilutedB Undiluted Emu sperm stored at 39°C 3–6D Dye exclusion (N-E) 3 Dye exclusion (N-E) DilutedF Ostrich sperm stored at 5°CE 48 Dye exclusion (N-E) ADye Australian Journal of Experimental Agriculture exclusion test results (Malecki and Martin 2000); swelling and membrane hydrolysis test results (Malecki and Martin 2005b). BPoultry diluents (BPSE, Lake’s). CEmu diluents (Modified Lake’s diluent). DDiluent dependent (Lake’s and NaCl-TES better than BPSE, Phosphate). EI. A. Malecki, P. K. Rybnik, A. Kowalczyk, E. Lukaszewicz and H. Naranowicz (unpubl. data). FEK diluent (Lukaszewicz 2002). Table 3. Viability of frozen-thawed emu and ostrich spermatozoa Freezing method Straws + SlowA Straws + SlowB Straws + RapidC Droplets + RapidD Droplets + RapidE CPA Viable sperm Sperm quality test (%) Emu spermatozoa 9% DMA 45 Dye exclusion (N-E) 9% DMA 10 Swelling (HOS) 6F Membrane hydrolysis (S-E) 9% DMA 10 Swelling (HOS) 3F Membrane hydrolysis (S-E) 6% DMA 15F Membrane hydrolysis (S-E) 24 Swelling (HOS) Ostrich spermatozoa 9% DMA 20 Dye exclusion (N-E) AIrrespective of the freezing rate, 1–7°C/min from +5 to –35°C and thawing in iced water bath (Malecki and Martin 2000). BFreezing rate 1°C/min from +5 to –35°C after cooling to +5°C and 15 min equilibration with DMA and thawing in iced water bath at ~5°C (Malecki and Martin 2005b). CDirect plunging into liquid nitrogen after cooling to +5°C and 15 min equilibration with DMA and thawing in iced water bath at ~5°C (Malecki and Martin 2005b). DDirect plunging into liquid nitrogen after cooling to +5°C and 15 min equilibration with DMA and thawing in hot water bath at ~60°C (Malecki and Martin 2005b). EDirect plunging into liquid nitrogen after cooling to +5°C and 15 min equilibration with DMA and thawing in hot water bath at ~60°C (I. A. Malecki, P. K. Rybnik, A. Kowalczyk, E. Lukaszewicz and H. Naranowicz, unpubl. data). FAs proportion of holes made by fresh untreated sperm. 1290 Australian Journal of Experimental Agriculture I. A. Malecki et al. would be preferred to 5°C. The reverse might apply to the ostrich, but more work is needed to optimise diluents and storage temperature. achieved, the ratite industries need to adopt AI technology and this can be accelerated by further development with support from the industry and government agencies. Semen freezing Freezing of emu and ostrich semen in liquid nitrogen (LN2) has been attempted using membrane-permeating cryoprotective agents (CPAs) such as glycerol, dimethyl acetamine (DMA) and dimethylsulfoxide (DMSO) at the final concentrations of 6, 9 and 12%, but frozen-thawed spermatozoa have only been evaluated in vitro (Malecki and Martin 2000, 2005b; I. A. Malecki, P. K. Rybnik, A. Kowalczyk, E. Lukaszewicz and H. Naranowicz, unpubl. data). Of the CPAs, DMA was found to be the most effective but its optimal concentration depends on the freezing and thawing method. In the emu, the loss of sperm function after freezing and thawing was considerable, as measured by sperm membrane integrity (HOS or N-E), fertilising ability (S-E) and motility (Table 3). Cryopreservation of emu semen in straws using a programmable freezer appeared to be as effective as direct plunging of diluted semen into LN2 when 9% DMA was used but, with direct plunging and rapid thawing, 6% DMA yields more viable spermatozoa than any other concentration. Motility of spermatozoa varied with freezing and thawing rates and the type and concentration of CPA: 40–60% of motile sperm were recovered following freezing at 1–7°C/min between +5° and –35°C or plunging into LN2 (Table 3). Thawing was carried out in an iced water bath at ~5°C. Similar motility results were also obtained following rapid freezing and thawing. Freezing of ostrich spermatozoa by direct plunging in LN2 reduced the number of viable spermatozoa by ~70%, as determined from N-E tests and proportions of motile sperm. It is clearly very early days and more research is needed before a viable protocol for freezing of emu and ostrich spermatozoa is developed and sperm banking becomes practical. References Conclusions The development of the AI technology for ratites has reached the stage of optimising methods for most efficient use of males for semen collection and protocols for semen holding and preservation. Females can be inseminated artificially with minimal discomfort, as they crouch voluntarily for this procedure. Fresh semen can be used for inseminations and possibly stored for a few hours as well, although this is yet to be determined. The underlying mechanisms controlling ovulation in emus and ostriches need to be understood if we are to achieve the most efficient male:female ratio. However, the two species provide us with different challenges, possibly because of differences in their mating systems. 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