AVIAN ARTIFICIAL INSEMINATION AND SEMEN PRESERVATION

George F. Gee

Patuxent Wildlife Research Center
Laurel, Maryland 20708
U.S.A.

Introduction
Many conservationists and aviculturists have joined in an effort to propagate non-domestic birds in captivity (Martin 1975). Intensive propagation techniques, some common to the domestic poultry industry and others peculiar to aviculture are used. Artificial insemination (Al) is one of these techniques (Gee and Temple 1978) that has been used extensively with domestic and nondomestic species. Al has been used successfully with cassowaries (C. Pickett and J. Mollen, pers. comm.), cranes (Gee 1969, Archibald 1974, Gee and Temple 1978), curassows and wild turkeys (G. A. Greenwell, C. Emerick, and P. Cohen, pers. comm.), ducks and geese (Johnson 1954, Watanabe 1957, Kinney and Burger 1960, Lake 1962, Pingel 1972, Skinner 1974, Kurbatov et al. 1976, and Tan 1980), eagles (Hammerstrom 1970, and Grier 1973), falcons (Bird et al. 1976, Boyd et al. 1977, Boyd 1978, and Lindberg 1977), hawks (Berry 1972, Temple 1972, and Corten 1973), peafowl (A. T. Leighton, pers. comm.), Pheasant and quail (Cain 1978, Gudelman et al. 1977, Wentworth and Mellen 1963), pigeons and doves (Owen 1941, and Szumowski et al. 1976), and psittacines (Harrison 1982).

The cryopreservation of avian semen is another new propagation technique that has been successful in chickens (Sexton 1979) and recently on a limited basis in propagating cranes (Gee and Secton 1979) and ducks (Watanabe et al. 1981). The U.S. Fish and Wildlife Service, the U.S. Department of Agriculture, and Agricultural Research Council’s Poultry Research Centre, Edinburgh, (Scotland) have active research programs in this area while several zoos, universities, and the International Crane Foundation have begun efforts to freeze and store avian semen.

AI as a Propagation Tool
AI programs have developed in response to a variety of needs in avian propagation (Smyth 1968, Martin 1975, Cooper 1977, Gee and Temple 1978). The most obvious need was to reduce or eliminate infertility (Szumowski et al. 1976, Lake 1978, Sexton 1979). In some mated pairs natural copulation can be difficult because of differences in body size, injury, or deformity, and in some, natural copulation maybe inhibited by behavioral difficulties. In other situations some females may be kept in separate pens because of incompatibility or the lack of a mate. Occasionally a productive female may be in a distant location separate from the male, where transfer of semen is the only alternative to infertility. Also, poor fertility in a mated pair can be improved through insemination with semen from another male.

AI is useful in carrying out the goals of special breeding programs for captive propagation. The genetic influence of one male in a population can be increased by using his semen to sire young from several females each season. Semen from several males can be used at one time to increase fertility or semen donors can be interchanged during a season so that chicks from the same female could have different sires. Hybridization between incompatible species is possible with Al and has been used extensively in poultry and waterfowl. Certain information can be gathered more quickly using insemination. For instance, a male’s potential for producing superior progeny or his potential fertility with several females can be determined more quickly through Al than with natural matings.

Semen collected has other uses. Laboratory studies of semen can be used to evaluate its reproductive potential (Sharlin et al. 1979), to evaluate semen diluents (Sexton 1977), to detect disease (Thurston et al. 1975, Stipkovits et al. 1978 and 1979, Ferrier et al. 1982) and to separate species and subspecies (Sharlin et al. 1979, Russman and Harrison 1982). Semen can be used to evaluate sperm preservation techniques and kept in the frozen state indefinitely (Sexton and Gee 1978, Watanabe and Terada 1976 and 1980, Watanabe et al. 1981).

Reproductive Anatomy and Physiology
To use Al, it is helpful to understand avian reproduction and to avoid confusion between bird and mammal reproductive systems. Male birds lack the accessory reproductive organs typical in mammals (Marshall 1961). The paired testes are located deep in the body cavity, above the abdominal air sacs and ventral to the cephalic end of the kidneys (Sturkie 1965). The testes consist of seminiferous tubules, rete tubules, and vas defferentia, but no septa or lobules. The sperm are taken from the simple epididymis on the caudal wall of the testis to the cloaca by the vas deferens (Sturkie 1965). The vasa deferens terminate as erectile teats in the urodeum (Sturkie 1965) and in some small birds, form a coiled structure above the dorsal lip of the cloaca, the cloaca) protuberance (Howell and Bartholomew 1952, Salt 1954, Wolfson 1952 and 1960, Middleton 1974). A few avian species have a copulatory organ for delivery of semen into the female’s oviduct (Bump 1969, Skinner 1974, Fujihara et al. 1976). Semen contains fluids secreted from the seminiferous tubules, epithelial cells of the reproductive tract, lymph from the lymph folds and erectile tissues in the cloaca, and sperm (Mann 1964, Lake 1966, Buxton and Orcutt 1975, Nishiyama et al. 1976, Servouse et al. 1976, Burt and Chalovich 1978, Gasparska et al. 1981).

Birds, the only class of vertebrates that consists exclusively of oviparous forms (Marshall 1961), lay eggs, generally within a day or two of ovulation. Although a few species have ovaries and oviducts on both sides, generally only the left side is functional (Sturkie 1965). The oviduct consists of an infundibular region, magnum, isthmus, uterus (shell gland), and vagina. The infundibular region receives the egg from the ovary and is the site of fertilization (Olsen 1942). The vagina is the passageway for the egg from the uterus to the cloaca and for the semen into the oviduct (Sturkie 1965). Sperm storage sites (sperm host glands) are present in the infundibulum and the utero-vaginal (UV) juncture (Bohr et al. 1962) and the UV-sperm host glands enable birds to lay several fertile eggs following a single copulation (Smyth, 1968).

Sperm host glands have been identified in ducks (Pal 1977), quail (Renden et al. 1981), and cranes (B. C. Wentworth, pers. comm.) as well as turkeys (Lorenz 1970) and chickens, and may be common to all birds. Sperm are released from the host glands on a continuous basis (Berke and Ogasawara 1969, Compton et al. 1977 and 1978, Compton and Van Krey 1979, Bakst 1980 and 1981). Although the release of spermatozoa from the uterovaginal sperm host glands at the time of ovulation or oviposition has been postulated, available information does not support the concept. Bakst (1980) reports that sperm numbers are less in oviducts with an ovum and may indicate that the lumina) spermatozoa are sequestered by egg formation.

Collection and Insemination
Al techniques are often grouped into cooperative or massage categories reflecting the different degrees of cooperation by male and female birds toward the human handler during semen collection and insemination. Cooperative Al requires a great deal of cooperation from the bird, massage Al is more successful with cooperation, and electroejaculation is successful without active cooperation from the bird. Cooperative semen collection and insemination was pioneered with sexually imprinted birds of prey being used in falconry (Hammerstrom 1970, Temple 1972, Berry 1972, Grier 1973). Falconers’ birds were already imprinted on their handlers and were encouraged to develop a sexual as well as social bond. The unrestrained male is encouraged to copulate, deposit semen in or on a suitable receptacle (Boyd and Boyd 1976, Boyd 1978). The semen, usually less than 0.1 ml, is aspirated into a suitable syringe or pipette to protect it from dehydration and contamination. The females are encouraged to respond to their handlers by assuming copulatory positions. The semen is deposited in the cloaca or everted oviduct of the receptive birds (Temple 1972, Berry 1972, Grier 1973, Boyd et al. 1977). Cooperative semen collections and inseminations require opportune timing to obtain an adequate number of samples and to obtain fertile eggs. Methods that intercept semen during natural copulation with other birds or dummy mounting devices are variations of the cooperative collection technique (Smyth 1968, Tan 1980).

The massage collection technique (Quinn and Burrows 1936) has been used for decades with domestic poultry and more recently with nondomestic birds (Gee and Temple 1978). With this technique, the bird is restrained by as assistant and an operator collects the semen. The process takes 5 to 10 seconds. A common practice is to have an assistant hold and stimulate the bird by stroking the inner shanks and the ventral abdominal region. At the same time the operator stimulates the region around the tail, abdomen, and vent by stroking with the left hand, from the post-dorsal region of the back to the interpelvic tail region, and then to the postlateral region below the tail. Next the operator forces the tail back with the left hand and the abdominal and sternal regions are stroked from anterior to posterior with the right hand. Usually the cloaca will respond to stimulation by a partial eversion and occasionally ejaculation. The cloaca is grasped dorsally by the thumb and index finger of the left hand. A small glass collection device (4-5 cm in diameter) is held in the right hand for semen collection. The first drop of semen is collected on the lip of the glass. The final steps (abdominal and cloaca) massage) are repeated and the remaining semen expressed from the cloaca with the fingers on the left hand. There are numerous minor modifications to the message technique that allow one person to collect and to inseminate semen. The modification may be an altered position for holding or stimulating a bird or it may be the use of a restraining device. Restraining devices are also used in team operation and can greatly facilitate the process in turkeys and waterfowl (Wantanbe 1957, Smyth 1968).

Procedures used to stimulate the male are repeated to prepare the female for insemination. An assistant holds and stimulates the bird by stroking the shanks and abdomen. The inseminator strokes the region around the tail, abdomen, and vent. The female responds with a partial eversion of the oviduct (Figure 1). Next, the assistant applies a gentle but steady pressure to the abdomen and vent to complete the eversion process, exposing the vagina. The inseminator inserts the inseminating device (a syringe, straw, pipette, etc.) into the exposed vagina and the assistant releases the pressure being applied to the abdomen (Figure 2). The inseminating device is allowed to return with the vagina to the relaxed position, and the semen is deposited. A deep vaginal insemination is usually preferable (Lorenz 1969, Ogasawara and Fuqua 1972) since the storage site (sperm host glands) is located at the utero-vaginal juncture (Bohr et al. 1962); however, a moderate depth of insemination gives satisfactory results (Smyth 1968, Wentworth et al. 1975, Bird et al. 1976, Boyd et al. 1977). Because of the possibility of injury which could interfere with egg production and fertility (Ogasawara and Fuqua 1972, Wentworth et al. 1975), the inseminating device is not forced into the utero-vaginal juncture. Although it is not always possible because of various anatomical and handling problems, it is best to deposit semen in the oviduct. Inseminations should be more frequent and timed to coincide with oviposition when semen is deposited in the cloaca rather than the oviduct (Gee 1969, Temple 1972, Berry 1972, Grier 1973, Archibald 1974).

Electroejaculation has been used for years with domestic mammals to collect semen but is not a common method for collecting avian semen. Several investigators do prefer electroejaculation of ducks and geese to other methods of collection (Serebrovski and Sokolovskaja 1934, Watanabe 1957, Chelmonska and Geborska-Dymkowska 1980) and, if its reported advantages can be applied to other birds, it may become a more common form of collection. Both the cooperative and massage methods of semen collection require training and cooperation to obtain good semen samples, and this training takes time before semen collection can begin. Electroejaculation does not require training. Electroejaculation requires the application of an electrical current of up to 80 volts.A positive pole is attached to the skin in the sacral region and the negative pole in a basin of water. The bird’s bill is placed in the water and the current applied to cause spasms and ejaculation (Serebrovski and Sokolovskaja 1974). Watanabe (1957) attached one pole of the electroejaculator to the sacral region and the other probe (a blunt rod) was inserted into the vent. He applied 30 volts (0.06 to 0.08 amperes) for 3 seconds, rested the bird for 5 seconds and repeated the same process until the bird flapped reflexly and ejaculated. The process was repeated for up to 5 times to obtain ejaculation. Duck semen collected by electroejaculation contained a greater number of spermatozoa and was released in larger quantities than that collected by the conventional massage technique.

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Special AI Techniques
Semen collection techniques can be modified for each species to compensate for anatomical, physiological, or behavioral characteristics (Figure 3). Many species of non-domestic birds produce small quantities of semen (about 10 ul) and special care must be taken in collection and protection. The volume of semen collected and sperm concentration varies greatly between species (Table 1), but in many species, part or all of the time, some birds produce samples too small for insemination.

Small samples can be aspirated into a small pipette or collected on the edge of a slide (Howell and Bartholomew 1952, Smyth 1968, Lake 1978). Small concentrated semen samples dehydrate rapidly and must be protected by adding a drop of diluent when collected. Although ejaculates produced by some males may be small, several collections may be made in a week (McDaniel and Sexton 1977, Gee and Temple 1978) and used to inseminate several birds. I n the Japanese quail, it only takes one day for sperm to pass from the testis to the cloaca (Amir et al. 1973, Clulow and Jones 1982). Although semen can be collected daily from some males, too frequent collection can cause swelling and reddening due to irritation of the vent. With special care, we have been able to obtain semen samples 5 days per week from cranes during the entire reproductive season.

Anal secretions is most birds do not complicate semen collection although many birds have anal glands (Quay 1967). However, the Japanese quail cloaca) foam gland produces a mucoid secretion, a meringue-like froth, that can interfere with semen collection. The froth is removed by gently squeezing on either side of the vent and wiping it away with a soft cloth before collecting the semen (Wentworth and Mellon 1963). Semen may be difficult to handle when contaminated with the secretion and it may be detrimental to sperm survival. The ani (Crotophaga sp.) is another bird that produces a large amount of anal secretions. However, few ani are kept in captivity and a semen collection technique has not been reported. In some larger birds like the rhea, a waxy or greasy material is present in the vent. Excesses of this material should be removed to avoid contamination of the semen.

Song birds are difficult to stimulate by massage but most have a well developed cloaca) protuberance during the breeding season where the semen is stored (Howell and Bartholomew 1952, Salt 1954, Wolfson 1952 and 1960, Middleton 1974). Small quantities of semen can be obtained from many passerines by applying gentle pressure to the protuberance. Densely packed sperm in the tiny drop of semen are difficult to keep alive because of dehydration during aspiration. At Patuxent, adding a small amount of avian diluent on (not in) the tip of the micropipette to the semen in the cloaca before collecting it can help maintain sperm viability.

The copulatory organs (phallus) in waterfowl, ratites, and tinamous can interfere with semen collection. The phallus, coiled within the ventral cavity of the cloaca in waterfowl and tinamous, uncoils as it erects during sexual excitation and carries semen along a twisted groove that runs the length of the organ. Early in the massage collection process, the phallus is everted (Smyth 1968, Skinner 1974) and after ejaculation, semen is collected at the base and tip of the phallus with a small funnel, tube, or suction device (Figure 4). The copulatory organ is

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Table 1. Semen volume, sperm concentration, and duration
of fertility for a variety of birds
Fertility
Species Volume Concentration (days) References
Domestic
Canary 10 ul J. G. Griffith (pers. comm.)
Chicken 0.5-0.8 ml 3.5×109 10-13 Smyth (1968)
Duck 0.3 ml 8 Watanabe (1957)
Goose 10-800 ul 8-16 Gee and Temple (1978), Oliver (1971), Johnson (1954)
Japanese Quail 10 ul 5×109 4-5 Smyth (1968)
Pigeon 10-20 ul Owen (1941)
Ring-necked Pheasant 50-250 ul 10×109 11 Smyth (1968), Cain (1978)
Turkey 0.2-0.3 ml 8×109 45 Smyth (1968)
Nondomestic
American Kestrel 14 ul 8 Bird et al. (1976)
Brewer’s Blackbird = 10 ul + Wolfson (1960)
Eclectus Parrot 50-100 ul Gee and Beall (unpub.)
Goshawk 20-30 ul Berry (1972)
House Finch 10 ul + J. G. Griffith (pers. comm.)
Prairie Falcon 50-100 u I 0.02×109 6-8 Boyd (1978), Boyd et al. (1977)
Red-tailed Hawk 0.1 ml 6 Gee and Temple (1978)
Swamp Sparrow = 10 ul + Wolfson (1952)
Sandhill Crane 10-200 ul 0.3×109 10 Gee and Temple (1978), Putnam (1982)
Wood Thrush 10 ul + Wolfson (1960)
Wattled Cassowary 1-5 ml C. Pickett (pers. comm.)
English Sparrow = 10 ul + Gee (unpub.)
Seaside Sparrow = 10 ul + Gee (unpub.)
Golden Eagle 0.2 ml 9 Grier (1973), Grier et al. (1973)
Cockatiel G. Harrison (pers. comm.)
Budgerigar G. Harrison (pers. comm.)
+ greater than domestic chicken less than domestic chicken 1 # sperm/ml of semen
approximate

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very delicate and rough treatment should be avoided or injury may result (Smyth 1968). In species that yield small semen samples, careful manipulation of the vent permits collection of the semen before the phallus is everted. The everted phallus contains numerous crypts and crevices on the surface, and eversion should be avoided if possible since the small semen sample is difficult to collect if spread over this irregular surface.

Feces is another source of contamination often encountered in waterfowl that should be avoided. Although contaminated semen should be discarded, fertile egg production is possible if the semen is cleaned of the coarse contaminants and inseminated immediately. Cloaca) insemination may be the better insemination route to avoid contaminating the oviduct (Perek et al. 1969). The cloaca should not contain feces at the time of insemination. Defecation will occur soon after insemination when the cloaca is full and fecal bacteria can kill large numbers of sperm and reduce fertility. It is possible to reduce semen contamination by withholding feed and water for 6 to 8 hours before collection (Smyth 1968). Also, forcing the bird to move around before capture may induce defecation and increase the number of clean samples collected.

From birds that do not respond to conventional collection techniques, semen can be collected after natural copulation with other birds or dummiers. The semen is intercepted during copulation by a collection device. In chickens, semen has been collected after copulation from the cloaca of the female, from an artificial cloaca, or from a collector fastened to the male (Smyth 1968). Lamert and McKenzie (1940) describe and illustrate in detail a male collection device used in the chicken. Semen has been obtained after copulation from blackbirds (Howell and Bartholomew 1952), chickens (Smyth 1968), and ducks (Tan 1980).

Insemination methods, frequency, and volume inseminated should be adapted to the needs of each species. Deep vaginal inseminations with large numbers of sperm produce the best fertility. In some nondomestic birds, eversion of the oviduct may be difficult or cause too much stress, reducing or terminating production. In some, the oviduct can be located by palpation and the inseminating device guided into the vagina to deposit the semen (Watanabe 1957) and others can be trained to open the cloaca in response to massage stimulation, exposing the oviduct and allowing vaginal insemination (Temple 1972). Satisfactory fertility can be obtained from semen deposited in the cloaca if inseminations are frequent (Gee 1969, Temple 1972, Berry 1972, Grier 1973, Archibald 1974).

Insemination frequency is dependent on other species specific characteristics such as sperm concentration and duration of fertility as well as the method of insemination. Birds with a short duration of fertility following insemination require more frequent inseminations than those with longer durations of fertility. Recommended insemination frequency varies from once every other day in Japanese quail (Lepore and Marks 1966) to once every other week in turkeys (Smyth 1968). The duration of fertility varies from 4 to 5 days for Japanese quail (Wentworth and Mellen 1963), and 6 to 8 days in some raptors (Grier 1973, Bird et al. 1976, Boyd et al. 1977) to 45 days in turkeys (Lorenz et al. 1959, Smyth 1968) (Table 1). Also, several repeated inseminations at the start of the insemination period improved subsequent fertility (Smyth 1968).

Semen volume inseminated depends on the sperm concentration and capacity of the reproductive tract to retain the semen. Often semen is of sufficient concentration and volume to inseminate several females from each ejaculation. Semen may be diluted for this and other reasons (Sexton 1976a). The volume of diluted semen inseminated may exceed the capacity of the bird’s oviduct to retain the semen and, although an adequate number of sperm are inseminated, fertility can be less than expected. The number of sperm needed per insemination in nondomestic birds is unknown, but in chickens and turkeys, the number has been estimated between 80 and 100 million spermatozoa (Sexton 1977b, Lake and Stewart 1978). More frequent inseminations may be necessary when sperm numbers per insemination are marginal (Meyer et al. 1980). For instance, a single sandhill crane ejaculate (200-300 million sperm/ml.; 0.05 ml/ejaculate – Gee and Temple 1978) may not be contain enough sperm to produce a satisfactory fertility rate. Based on more than 10 years of experience with cranes, we recommend repeated insemination every other day thereafter, and inseminations after every oviposition to obtain satisfactory fertility.

Management
Al is only one of several methods used to correct infertility and all alternatives should be considered. AI is very time consuming and should be avoided if possible. Artificial insemination of nondomestic birds is labor-intensive since 2 persons are often necessary to collect and inseminate, and an additional person is often needed to help capture or separate birds (Gee and Temple 1978). Since natural copulation in properly mated birds generally results in better fertility than from artificially inseminated birds, a change of mates may improve fertility more than Al. However, Al can be as effective as natural coatings in the crane (Gee 1969), American kestrel (Bird et al. 1976), and chicken (Smyth 1968), and is more effective in the domestic turkey (Smyth 1968). Also, all eggs that fail to hatch need to be examined closely to determine fertility. A clear egg is not necessarily infertile. Improper incubation conditions such as overheating or cold, bacterial contamination, nutritional deficiencies, disease, and others can destroy the early embryo (Taylor 1949). Obviously, Al would not be a solution to such problems.

Proper physical, physiological, nutritional, and behavioral conditions must be provided to obtain reproduction. Of the environmental conditions, light, temperature, and humidity are three of the most powerful factors influencing reproduction. For a more detailed review of avian breeding seasons and proximate and ultimate factors, see Lack 1950, Moreau 1950, Lofts and Murton 1968 and 1973, Immelmann 1971, Seventry 1971, Farner 1973, and Murton 1978. In most birds, semen production begins before and continues until afterthe end of egg production. However, asynchrony, the production of semen and eggs at different times in the reproductive season, does occur and can be corrected by manipulation of day length in some species. For example, turkey toms can be exposed to long light days several weeks before the hens to guarantee good semen production before the first eggs are laid (Nestor and Brown 1971). Many birds will not produce eggs or semen if the length of day is too long (Murton 1978). Like light, improper temperature and humidity can terminate or delay reproduction, especially in desert birds (Seventry 1971, Immelman 1971). Building and nest design, sickness, inadequate diets, and aging disabilities are other factors that can affect fertility rate.

Late in the season, semen production may cease before egg production is completed. However, in the chicken and turkey, most semen from older males or from males late in their reproductive cycle produces satisfactory fertility (Woodard et al. 1976, Ansah et al. 1980). An immune response against sperm may develop late in the reproductive season and reduce fertility (Yu and Burke 1979, McCorkle et al. 1981).

When circumstances indicate AI is necessary for increasing fertility or to complete special breeding programs, the birds should be trained to accept the technique. Behavioral accommodations are of great importance in artificial insemination of nondomestic birds, especial those taken from the wild. Stress is difficult to avoid in artificial insemination of nondomestic birds, but using the same team, training birds to accept artificial insemination procedures, and avoiding injury to the birds when handling all reduce stress. Disturbance can be further reduced by restricting visitation to the non-breeding season, and conducting all chores on a constant time table.

The training process in a few birds may upset the female rather than calm her down and if continued, may interfere with the onset of egg production. The training period in these cases should be terminated with both male and female. Insemination can be reinstated when egg production starts and the bird is more receptive to stimulation. Semen volume and response to semen collection often can be improved by placing a bird near other birds. Visual and auditory displays by other reproductively active birds may stimulate reproductive activity in surrounding pairs and even in single birds, and it can strengthen pair bonds (Wickler 1980).

Little can be done to increase semen yield from a content and healthy bird, but several steps can be taken to protect the semen collected and make the best use of it in insemination. A water bath or insulated container reduces temperature fluctuations and a closed tube reduces dehydration and contamination. A diluent increases semen volume, reduces the risk of dehydration, and if sperm concentration is adequate, makes it possible to inseminate several birds from each ejaculate. Diluent reduces sperm concentration, bacterial contamination, and controls pH and osmolality. All the tubes and inseminating devices that come in contact with the semen should be clean. Since detergents can be especially harmful to semen samples, all equipment and supplies should be thoroughly rinsed with clean water before use. For reviews of factors detrimental to sperm survival, see Mann (1964), Lake and Steward (1978), Lake (1969), and Smyth (1968).

Semen Evaluation
Although the most reliable semen test is the production of fertile eggs, semen for use in Al can be evaluated in the pen when it is collected and later in the laboratory. The color of good semen when it is collected is characteristic for each species. Most good samples range from a light white (frosted glass appearance) in species with low sperm concentrations, to a white chalky or milky appearance in species with high sperm counts. Fecal contamination discolors the semen to shades of brown or green. In geese, a green color is common because of their large intake of grass during the breeding season. Occasionally, flecks of blood may be present resulting from excessive force during collection or injury (Smyth 1968). Samples that are consistently contaminated with feces may need to be diluted with antibiotics to reduce the loss of sperm and the anitibiotic, tobramycin, may even increase fertility when used as a diluent in “clean” semen (Sexton et al. 1980).

The consistency of a\ semen sample ranges from that of water to that of heavy cream in species with the more concentrated samples. Samples that appear to be sticky or stringy are often contaminated. Some of these semen samples begin as a clear fluid in the collecting device and turn white as the urates precipitate out. A watery semen may indicate an excessive amount of lymph (transparent fluid) in the sample. The use of excessive force on the cloaca during collection can cause a surplus of lymph in the semen. These watery fluids, like fecal and urate contaminants, adversely affect spermatozoa, especially if the semen is held for some time before insemination (Smyth 1968, Lake 1971, Fujihara and Nishiyama 1976).

Small representative portions, taken from the semen before insemination can be examined in the laboratory for sperm number, motility, morphology, and larger samples for metabolic rate and semen composition. Perhaps the simplest measures of semen quality are sperm number and motility. Sperm number can be estimated from a semen score for density, spermatocrit, counting in a hemocytometer or in an automated density, spermatocrit, counting in a hemocytometeror in an automated counter. Sperm concentration can be evalutated in semen mounted on a hanging drop slide, under a cover slip on a slide, or in a capillary tube. This is a useful way to distinguish semen quality in species with low sperm concentrations like cranes (Putnam 1982); however, semen from species with higher sperm concentrations first require dilution. The scores can be calibrated by comparing them to actual sperm counts. The spermatocrit, a simple measure of sperm concentration, is useful in characterizing semen produced in quantity (>0.1 ml) and containing a large number of sperm per ml (>3×109) (Arscott and Kuhns 1969). The semen sample is loaded into the standard microhematocrit capillaries and centrifuged. The percent by volume is determined in several ways. The semen is diluted (if necessary), fixed, and the sperm counted in a hemocytometer or in an automated counter(Jones and Wilson, 1967). Optical density of a diluted semen sample can be measured and sperm number determined from a previously established standard curve (Kosin and Wheeler 1956, Carson et al. 1955).A reasonable estimate of sperm concentration is important in determining if the insemination dose will contain an adequate number of sperm.

Sperm progressive motility is an estimate, based on a scale from 0 to 100, of the percentage of spermatozoa moving in a forward motion. This is another quick indicator of semen quality. A drop of semen is placed on a clean microscope slide under a cover slip and several areas examined at a 430 magnification. Since some live cells are inactive, it is not an estimate of all live cells.

Examining sperm morphology using conventional miscroscopes can provide information of the percentage of live cells in the semen, the percentage of abnormalities, and the size of cells. One of the easiest determinations, a live-dead count from an eosin-nigrosin stained slide, makes it possible to evaluate the number of live sperm inseminated (Gee and Sexton 1979). It is a more time consuming determination than progressive motility, but it can be determined in the laboratory long after the insemination, usually without a significant loss in accuracy. However, there are confounding factors such as excessive moisture in the atmosphere that can make the staining process less definitive (Ogasawara et al. 1976). Abnormalities can be determined from a variety of preparations including the eosin-nigrosin stained slide. Good slide staining techniques aid in delineating parts of the spermatozoa such as the head from the acrosomal cap and mid-piece (Sharlin et al. 1979). Abnormalities in sperm can be used to evaluate semen from individual males and to determine effects of diluents and storage. Sperm head size determined from properly stained slides can be used to distinguish between subspecies (Sharlin et al. 1979, Russman and Harrison 1982) and to predict fecundity within subspecies (Sharlin et al. 1979). Electron microscopy is useful in detecting membrane and fine structure abnormalities in spermatozoa, but its usefulness is generally confined to research.

Metabolic rate in semen samples can be determined using a variety of tests such as methlene blue reduction and oxygen consumption (Smyth 1968). The effects of various diluents, environmental factors, and individual differences can be evaluated. Determinations require precise laboratory control and may require very specialized equipment.

The chemical composition of semen from most avian nondomestic species is unknown and even in the domestic fowl it is not very useful in distinguishing between good and bad semen samples (Mann 1964, Burt and Chalovich 1978). Semen pH and osmolality are two factors that vary from species to species and should be considered when diluting semen. In our laboratory, we have recorded semen pH in a range from 6.0 for a duck to 8.0 for a crane. Laboratory tests can be used to evaluate differences between new diluents or techniques, but eventually the semen must be inseminated and fertile eggs produced to prove its value. Satisfactory fertility has been obtained from semen that scored poorly in laboratory tests, especially frozen-thawed semen (Sexton 1976b). Cryoprotectants and freezing can affect sperm motility and morphology without destroying the ability of the frozen-thawed semen to produce fertile eggs.

Equipment and Supplies
The basic equipment used to collect and inseminate semen is relatively simple and inexpensive (Corten 1973). Semen collecting devices include those used to hold the bird (cones, stands, and jackets) and those used to catch the semen (cups, funnels, test tubes, capillary tubes, pipettes) (Smyth 1968). Electroejaculation requires special equipment and devices capable of delivering measured electrical charges and appropriate probes adaptable to the species. Containers of semen can be held in a small rack and case until they are needed for insemination. If the sample is held for more than a few minutes, the case provides protection from temperature shock, direct sunlight, and contamination. A thermometer is useful in determining the case temperature.

Inseminating equipment includes syringes, pipettes, straws, or eye droppers and devices to hold or add diluents. A flexible tube attached to a mouthpiece at one end and a semen straw at the other is a useful method for inseminating large numbers of birds (Smyth 1968).

The animal facilities should be free of obstructions to reduce the chances of injury and to accommodate a quick, less stressful capture. Also, clean facilites reduce the risk of semen contamination and soiling of the bird during capture.

A few pieces of laboratory equipment like the microscope are all that are needed for the routine examination of semen. Progressive motility estimates require only a clean slide and coverslip, but a hanging drop slide is useful for lengthy microscopic studies of living samples. A variety of stains are needed for live-dead counts and to perform special morphological studies. A balance is needed to weigh out chemicals, and to prepare stains and other supplies. Other supporting pieces of equipment and supplies include cell counters, slide trays, record books, and photographic attachments. Semen and diluent pH determinations require pH paper or a pH meter. Osmolality can be determined from small samples using a vapor point osmometer. Tests of semen metabolic activity may require respirometers or spectrophotometers.

Storage and Preservation
Although the best fertility rates are obtained from inseminating semen immediately following collection, fresh semen can be stored for several hours or more (more than a day in chickens) without destroying fertility. A rate of thirty-seven percent fertility after 36-37 hours storage is possible in chickens (Lake et al. 1959). Semen storage for an hour or more requires temperature control and protection from contamination and drying. Sperm of most species’ survive best at near freezing temperatures (Gee and Temple 1978, Sexton 1979). Since bacterial contamination can cause a rapid destruction of sperm cells in storage, sources of contamination must be avoided. Diluents can be used to introduce antibacterial agents, stablize pH and osmolality, and in other ways extend the viable life of a semen sample (Smyth 1968, Gee and Temple 1978, Sexton et al. 1980).

Semen preservation in the frozen state has been demonstrated in a few birds and makes it possible to store semen for many years (Sexton 1979, Gee and Sexton 1979, Watanabe and Terada 1980, Watanabe et al. 1981). Fertility rates with frozen-thawed semen can be improved using intrauterine (Watanabe and Terada 1976, Marquez and Ogaswara 1977) and intraperitoneal (Harris 1968) inseminations, but good fertility has been obtained using the conventional methods described earlier (Sexton 1976).

There are four critical steps in cryopreservation of semen: pre-freeze, freeze, storage, and thawing (for a recent review see Mazur 1980). Important pre-freeze considerations are time held, dilution rate, contamination, diluent, and pH of the semen. The time a sample should be held before it is frozen depends on the conditions of storage and are species specific (Sexton and Gee 1978). The number of good quality semen samples (90% or more of live motile sperm) decrease with time held before freezing. However, if the samples that deteriorate are discarded and only good samples are frozen, fertility rates from Al with frozen-thawed semen held before freezing are similar to those frozen soon after collection.

cryoprotectant, cryoprotectant level, equilibration time, and freeze rates are the most critical variables in the successful preservation of semen. The cryoprotectant level required to provide protection against freeze damage is dependent on the diluent, pH, equilibration time, and varies between species (Watson 1978). Of the two most common cryoprotectants, glycerol and dimethylsulphoxide (DMSO), only glycerol has to be removed before insemination since it has an antifertility effect in birds (Nevelle et al. 1971, Sexton, 1979, Lake et al. 1980). Levels of glycerol that provide good protection against freeze damage arange from 8% to 15% (Brown and Harris (1963, Neville et al. 1971), and for DMSO, from 4% to 10% (Sexton 1979, Gee unpublished). Freeze rates are different for each cryoprotectant as are the mechanics for sample freezing. Glycerol preserved samples can be frozen in a variety of successful schemes (Hawk 1979). DMSO rate seems to require a precise freeze rate of VC per minute from 5°C to –20°C, 50°C per minute from –20° C to –80°C, and 160°C per minute from –80°C to –196°C (Sexton 1980). New successful DMSO schemes may be possible when a greater number of experimenters adopt the use of DMSO.

Storage and thawing conditions are important in the recovery of viable sperm from frozen-thawed semen. Deterioration of the sample is prevented if it is stored at liquid nitrogen temperatures (-196° C), but some deterioration has been recorded at higher temperatures (-80° C) (Graham 1975). Most techniques thaw samples at room temperature or in a crushed ice bath; however, the effect of thawing rates on sperm survival needs more study (Gee and Sexton 1979).

Frozen-thawed semen should be inseminated by the most effective route and at the most advantageous time during the reproductive cycles. Frozen-thawed semen has less energy reserve that a fresh sample due to storage while preparing it for freezing and the cells may be less active due to this and inhibition as soon as possible after it is prepared for insemination and all adverse environmental conditions avoided.

Summary
Artificial insemination is a practical propagation tool that has been successful with a variety of birds. Cooperative, massage, and electroejaculation and modifications of these three basic methods of semen collection are described fora variety of birds. Semen color and consistency and sperm number, motility, and morphology, as discussed, are useful indicators of semen quality, but the most reliable test of semen quality is the production of fertile eggs. Successful cryogenic preservation of avian semen with DMSO or glycerol as the cryoprotectant has been possible. Although the methods for preservation require special equipment, use of frozen semen requires only simple insemination supplies.

Acknowledgements
The author acknowledges the support of many individuals throughout the U.S.A. of our efforts in artificial insemination of nondomestic birds. A very special thanks to my colleagues over the years at the Patuxent Wildlife Research Center who have made artificial insemination and semen preservation possible in cranes.

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