Abstract

The various subspecies of hartebeest (Alcelaphus buselaphus) formerly ranged throughout all Africa. Encroachment of farming and livestock has resulted in a critical loss of habitat and food and water resources for the hartebeest. These factors, along with hunting for meat and sport, have taken their toll on hartebeest populations in the wild.³ The subspecies A. b. tora and A. b. swaynei occurred formerly from Southern Egypt to Somalia, are classified as endangered by the IUCN/SSC.³ Much of the remaining range of A. b. tora, in southern Sudan and northwestern Ethiopia was devastated by drought in the early 1980s, and only a few animals are thought to survive. Alcelaphus buselaphus has also disappeared from most of South Africa but is still locally common in parts of Botswana, Namibia, Tanzania, and Kenya.⁴

Successful captive propagation of hartebeest in zoologic institutions is an important step in the conservation of this species. This need will become more critical, as habitat becomes increasingly threatened and hartebeest numbers subsequently decline. The captive management of hartebeest in zoologic institutions has proven challenging. At present, three subspecies exist in six North American zoologic institutions. The Wildlife Conservation Society’s St. Catherines Island Wildlife Survival Center (WSC) houses two of these subspecies: the Jackson’s hartebeest (A. b. jacksoni) and the Cape hartebeest (A. b. caama). The WSC has been working with the Jackson’s hartebeest since 1978, and has been very successful at maintaining breeding herds. The WSC now holds all the remaining Cape hartebeest in North America.

The purpose of this study was to perfect semen collection, evaluation, and long-term storage techniques for these two subspecies of hartebeest. We are currently evaluating whether there are any seasonal differences in the sperm quality and quantity in these animals. The long-term objective will be to develop assisted reproductive techniques (e.g., artificial insemination) in captive hartebeest. This technology will allow the capability of importing semen into this country from captive and wild hartebeest in Africa and Europe to increase the genetic diversity of current captive populations.

Adult male hartebeest were immobilized in individual holding stalls. Carfentanil (Wildnil, Wildlife Laboratories, Inc, Fort Collins, CO) at 2.1–2.4 mg per animal was the primary immobilizing agent used in this study. Once the animals were sternal they were given 10 mg xylazine (X-ject SA, Burns Veterinary Supply, Inc., Rockville Centre, NY) intravenously incrementally as needed to provide muscle relaxation and analgesia. An intravenous catheter was placed in the lateral saphenous vein for fluid and drug administration throughout the procedure. Anesthesia was monitored via pulse oximetry, respiration, and digital pulse rate.

Once anesthetized, semen was collected from hartebeest by electroejaculation, using a 38 mm diameter probe with longitudinal electrodes. A standardized electroejaculatory regimen consisting of administering electric stimuli in sets of 10 at 2, 3, and 4 volts (series 1), 3, 4, and 5 volts (series 2) and 5 and 6 volts (series 3) were used to collect semen.²

Laboratory calipers were used to measure the width (W) and length (L) of each testicle, recorded in cm. Combined testes volume is calculated by using the following formula (W² × L) × 0.524 for each testis.¹ Representative ejaculate and sperm traits are shown in Table 1.

Table 1. Ejaculate and sperm traits of Jackson’s and Cape hartebeest

The semen was evaluated grossly and noted to have a distinct pink color. Examination of the ejaculate showed that no red blood cells were present and the preliminary diagnosis is that the semen contains porphyrins. Preliminary results indicate that the presence of porphyrins is not detrimental to short-term sperm storage. Semen pH, volume, and sperm concentration were measured. Percent sperm motility and progressive motility (forward progressive movement on a graded scale: 0 = no movement to 5 = rapid, steady forward progression) were assessed by examining 5 µl of raw ejaculate at 37°C. A 20 µl aliquot of the raw ejaculate was fixed in 0.3% glutaraldehyde and sperm morphology was assessed by phase contrast microscopy (×100). Sperm morphology assessments were conducted by classifying 100 spermatozoa as normal or abnormal. In the Jackson’s hartebeest, one of the most striking findings is significant abnormal sperm morphology. The abnormal classes of sperm identified are: cytoplasmic droplet, bent midpiece with droplet, abnormal midpiece, coiled flagellum, bent flagellum with a cytoplasmic droplet, bent flagellum without a droplet, bent mid-piece without a droplet, biflagellate, and other (includes spermatids; Table 1). In contrast, Cape hartebeest have high proportions of morphologically normal spermatozoa (Table 1). Table 1 presents the results of the fresh semen evaluation for Cape and Jackson’s hartebeest.

The semen was divided into three aliquots and extended in either TEST (0.4% glucose, 4.83% tes-n-tris, and 1.15% tris), BF5F (1.6% glucose, 1.6% fructose, 1.2% tes-n-tris, 0.2% tris and 0.5% sodium triethanolamine lauryl sulphate) or Tris diluent containing 5% glycerol to a concentration of ~200 ×10⁶/ml.¹ Extended semen was assessed for motility and status and cooled to 4°C over 2 hours. Cooled extended semen was loaded into 0.25 ml straws and frozen over liquid nitrogen (LN2) for 8 minutes before plunging into LN2. Straws were stored in LN2 for a minimum of 24 hours before thawing. Straws were thawed by immersion in a water bath at 37°C for 15 seconds. Straw contents were released into sterile microcentrifuge tubes and assessed for motility, status, and viability at 37°C. Thawed sperm was diluted 1:3 with Talp medium supplemented with pyruvate and 5% fetal calf serum. Talp was added in increments to reduce the osmotic shock of removing glycerol. After 30 minutes, diluted sperm was re-assessed for motility, status, and viability. Assessments were repeated hourly. Table 2 presents the preliminary results of cryoprotectant addition, cooling, and thawing.

Table 2. Cape and Jackson’s hartebeest sperm characteristics after cooling, freezing, and thawing in different extenders

The results of this study demonstrate that hartebeest spermatozoa do not respond well to the cryopreservation process and low numbers of motile sperm are recovered. The higher pre-freeze sperm motility in the Cape hartebeest resulted in higher post-thaw values. However, preliminary results indicate that proportionally, similar numbers of both Jackson’s and Cape hartebeest sperm survived the freeze-thaw process. BF5F results in higher pre-freeze and post-thaw motilities, but insufficient data is available to compare this statistically. Further studies will investigate whether the high numbers of abnormal sperm are related to possible seasonality of the species or to a lack of genetic diversity in the captive population. Also, the effects of increasing glycerol concentration will be investigated to attempt to increase the numbers of post-thaw motile spermatozoa.

Literature Cited

1.  Harrison RM, Dominique GJ, Heidger PM, Roberts JA, Schlegel JU. Vasectomy in the Rhesus monkeys. Surgical techniques and gross observations. Urol. 1977;9:639–644.

2.  Howard JG, Bush M, Wildt DE. Semen collection, analysis, and cryopreservation in non-domestic mammals. In: Morrow DA, Saunders WB, eds. Philadelphia, PA: Current Therapy in Theriogenology. 1986:1047–1053.

3.  Kingdon J. The Kingdon Field Guide to African Mammals. San Diego, CA: Academic Press; 1997.

4.  Nowak RM. Walker’s Mammals of the World, Fifth ed. Volume II. Baltimore, MD: The Johns Hopkins Univ Press; 1991:1450–1451.