ARTIFICIAL INSEMINATION IN POULTRY

Artificial insemination (AI) is a novel tool of assisted reproductive technology, which is being successful in improving fertility in most of the domestic animals including poultry. The profitability of any poultry breeder operation depends on the number of salable chicks produced in the hatchery, which again depends on the fertility of the breeder flocks. Fertility is a complex component that depends on factors like genetics, nutrition, and management of both sire and dam. Today, poultry are intensively bred for meat purpose, which might have negative influence on the fertility as body weight and fertility are negatively correlated. AI is of very useful when there exist a wider weight difference between sire and dam where mating will be difficult. This is true especially with dwarf broiler breeders and turkeys. Further, AI allows large number of females to be inseminated from small number of elite males allowing quick genetic improvement with reduced maintenance costs.

Selective mating, poor fertility due to sexing problems, seasonality of breeding can also be overcome by AI technology. Hence, AI is a promising tool for improving the fertility in commercial poultry breeding operations. The two major steps in artificial insemination include the collection of semen from the males and deposition of semen in the female oviduct. In addition to the above, the current paper focus on the preparation and training of males for semen collection, various semen diluents, semen quality assays and monitoring fertility in artificial inseminated flocks. AI is commonly practiced in turkey and broiler breeders while the technique was also successful in Guinea Fowls, Japanese Quails, Emu and Ostrich.

 Preparation of males for semen collection

The breeder males, which are selected as sires for next generation, should be separated from other female birds and housed separately to prevent natural mating. Broiler breeder males, guinea fowls, and Quails can be individually caged which allows easy handling. The birds should be fed with appropriate breeder male rations and should be given a period of about one week to accustom in the new housing/cages. No change of care taker/ handler is entertained as it will lead to handling stress or panic in bird thereby causes loss in semen quality. The feathers around the soft part of the abdomen should be trimmed of for easy and clean semen collection

Semen collection

Semen should be collected during early hours of the day, twice a week for optimum semen quality. Four methods of semen collection were reported to be successful in different poultry species. The widely accepted, non invasive and most common method of semen collection is the abdominal massage technique. The birds are to be trained for semen collection for a minimum period of one week. However, guinea fowls requires longer training period of about three weeks  as these birds are timid and takes longer time to accustom with the handler. The sides of the soft part of the abdomen and the bilateral region of the pygostyle were massaged rapidly and continuously until the bird responded by protruding the papillae from the cloaca. On the protrusion of papillae, the thumb and the index fingers of the hand were positioned conveniently to squeeze out the semen gently. Abdominal massage technique can be employed for semen collection in chicken, turkeys, guinea fowls, waterfowls, and quails.

The second method of semen collection is the electro ejaculation technique, which can be employed for semen collection in water fowls, psittacine birds and pigeons. A current of 5-30 volts can be applied using special probes in the dorsal body wall following anesthesia of the bird. Improper collection by this method may lead to serious burns or death of the bird the ejaculates are frequently contaminated with droppings.

Other methods of semen collection include the teaser female and the artificial cloacae technique, which are employed for semen collection in Emu and Ostrich. Here, the birds voluntarily copulate on special devices, which may be a dummy female, or ejaculate in an artificial cloaca in response to a behavioral stimulus like adequate vocalization or food transfer followed by a copulatory display.

Equipments

Artificial Insemination requires relatively simple equipments but the cleanliness of the equipments are of prime importance. Semen can be collected directly in a small glass funnel following abdominal massage technique. In certain birds/ species, where the volume of the semen is low, the drop of semen coming out of the papillae can be directly aspirated into a glass tuberculin syringe. Individual glass cannulas can be used for insemination, which prevents the spread of diseases to other birds. All the equipments should be sterile and dry, as water is spermicidal thereby causes fertility loss.

In case of Emu and Ostrich, the artificial cloaca can be designed using a PVC pipe of about 10 cm diameter and 30 cm length. A rubber lining of appropriate size should be used inside the tube with a collection cone and collection vial at one end of the pipe. The space between the PVC tube and the liner should be filled with warm water (42-45o C). The glass containers and collection vials used for handling semen may also be wrapped with aluminum foil, which prevents the passage of the light as light is detrimental to spermatozoa. The semen collected should be held at 18 to 20oC through the process of dilution and insemination.

Semen Dilution

The undiluted raw semen gradually loses its fertilizing capacity one hour after collection. Therefore, if raw semen is used for insemination, artificial insemination should be carried out within the above-specified period. This is impossible while inseminating large number of birds in a commercial setup. Further, raw semen with high spermatozoa concentration may be sticky where the sufficient volume of semen may not be deposited at the site of insemination. The use of semen extenders helps in maintaining the fertilizing ability of spermatozoa until about six hours of storage further increasing the volume, facilitating the insemination of large number of birds. The diluents should have an optimum nutrient environment, pH and osmolarity which helps in maintaining the morphology and livability of spermatozoa thereby maintaining the fertilizing capacity.

Semen diluents namely the Beltsville Poultry Semen Extender, Modified Beltsville Poultry Semen Extender, Lakes Semen Extender, IMV poultry semen preservation media etc, can be used. Normal Saline can also be used when insemination is carried out immediately after collection. The diluents can be added in the ratio of 1:2 or 1:3 based on the concentration of the spermatozoa in the semen. 

Semen Quality

Semen collection should be carried out with almost care, so as to collect it free from droppings or blood. Excessive pressure while collection may cause damage of small blood vessels thereby adding blood to the semen. Good quality Semen without contamination has a pearly white colour with thick consistency.

The other major semen quality parameters include the percent spermatozoa motility, per cent live spermatozoa and percent abnormal spermatozoa. The per cent spermatozoa motility is the percentage of spermatozoa that are motile on its own power. It determines the movement of spermatozoa from the site of insemination to the site of fertilization in the female oviduct. Motility can be accessed by placing a drop of semen on the microscopic slide with cover slip under a microscope. The quantum of spermatozoa having progressive forward motility can be expressed as percentage.

The percent live spermatozoa and abnormal spermatozoa can be accessed by eiosin- nigrosing staining procedure. With respect to livability the spermatozoa were classified as live (unstained cells) and dead (every cell stained by eosin). Spermatozoa within the fraction of live cells were classified as morphologically normal (normal head with well-marked acrosome and visible tail) or deformed (swollen head, bent neck, defective mid-piece, coiled tail, lack of tail etc.).

A minimum spermatozoa concentration of 1000 millions/ ml in raw semen is required for carrying out dilution and artificial insemination. Birds with lower spermatozoa concentration should be culled. The percent motility, live spermatozoa, and morphologically normal spermatozoa should be above 80 % for optimum fertility. The volume of semen and the concentration of spermatozoa in different species of poultry are given below. However, the semen quality varies between different breeds, strains, plane of nutrition, season etc.

SpeciesSemen Volume (ml)Spermatozoa Concentration (x106/ml)
Chicken0.30 – 0.703000 – 4000
Turkey0.12 – 0.221800 – 3500
Guinea Fowls0.04 – 0.101500 – 3500
Japanese Quails0.02 – 0.040500 – 0900
Duck0.75 – 1.000800 – 1500
Goose0.18 – 0.900230 – 2020
Emu0.10 – 1.301500 – 3500
Ostrich0.10 – 1.501500 – 4000

Artificial insemination

Artificial insemination should be done in the evening hours after oviposition, as presence of hard shelled eggs is known hinder the transport of spermatozoa in the oviduct. The volume to be inseminated, should be determined by the concentration of spermatozoa in the diluted semen. The usual dose is 100 million spermatozoa per insemination. Insemination can be done at lower doses when the frequency of insemination is increased. Insemination of 100 million spermatozoa once in a week, maintains optimum fertility in most of the poultry species.  

The bird to be inseminated should be held by an assistant. The oviduct can be everted by applying pressure over the soft abdominal region. Any adhering dirt or fecal particles should removed by cotton swabs. Glass tuberculin syringe/ cannula containing appropriate volume of semen should inserted to a desired depth, pressure should be released over the abdomen, and semen should be deposited gently without any disturbance to the bird. In case of Ratides, the voluntary crouch method of insemination can be practiced.

Monitoring fertility

Monitoring of AI flocks is of prime importance in poultry breeder operation. Traditional methods include checking for optimum semen quality, candling of hatching eggs, and break open analysis of unhatched eggs.

A recent and most effective method of monitoring fertility is the inner perivitelline spermatozoa penetration assay. The advantage of this method is that the fertility can be predicted on the particular day in which the egg was laid. Unsettable, misshapen and cracked eggs which are usually discarded can be subjected to this technique in large breeding flocks. Following insemination/ natural mating the spermatozoa deposited in the oviduct are stored in the sperm storage tubules which allows fertilization of successive eggs. As the bird lays eggs, the number of spermatozoa in the sperm storage tubules get depleted and the number of spermatozoa which interacts with the eggs decreases. The above technique counts the number of holes made by the spermatozoa in the perivitelline layer of the egg on its way to fertilize the female germ cell. It was reported that the chicken and turkey eggs have 50% probability of being fertile when around three spermatozoa penetrated the inner perivitelline layer over the germinal disc and showed maximum fertility when more than six spermatozoa penetrated this region.

IPVL Sperm penetration holes observed in the perivitelline layer of chicken egg.

To conclude, Semen quality is one the major determinant of fertility, which depends on the genetic, nutritional, and environmental factors. Standard breeder management coupled with proper semen handling yields semen of superior quality, which on insemination at optimum dose and frequency yields higher fertility. In addition, monitoring of fertility with modern techniques allows immediate action when low fertility was predicted. This increases the number of chicks per sire, enabling faster genetic gain in breeding operation and constant chick supply in a commercial setup.

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