The embryo transfer (ET) technique offers a powerful potential for genetic improvement in alpacas and llamas. The technique consists of collecting an embryo from a female donor through uterine lavage 7 to 8 days after mating. The embryo is then transferred to a synchronous recipient female to carry the pregnancy to term. As in other livestock species, embryo transfer offers several advantages, including the multiplication of females with high genetic merit and treatment of some aspects of infertility. It also permits a rapid repopulation of camelids as well as the preservation of endangered species. In addition, sires with high genetic value may be used throughout the year (not only during the breeding season) provided they are maintained in top condition with nutrition and health care.
Embryo transfer programs utilize a database with detailed records of the embryo’s pedigree. This information plays an important role in the detection of genetic flaws or hereditary defects that may be transmitted to subsequent generations. This information helps improve genetic selection.
Embryo transfer technology may be performed with or without hormonal superstimulation of the ovaries. A project conducted in Peru (Mega Alpaca Project, 2008) showed that it is possible to collect up to two embryos per donor per month without stimulation. This would result in potential production of 8 to 10 offspring per year instead of just one using natural techniques.
The potential for production of even higher number of embryos per year is possible through the use of multiple ovulation and embryo transfer (MOET). Protocols for superstimulation (superovulation) have been reported by several authors.1–4 Commercial alpaca embryo transfer in Australia reports an average of 2.5 to 3 embryos per uterine flush, with a potential for production of up to 21 embryos per individual per year.5 Pregnancy rates following transfer of embryos vary from 40% to 70%.5–7
Finally, the embryo transfer technique enables the preservation of endangered species such as vicunas, guanacos, and some breeds of alpacas and llamas through the use of interspecies embryo transfer.6,7 Cryopreservation of embryos could increase the application of this technology even more for genetics improvement and preservation of genetic diversity.
This chapter will review the history of embryo transfer in alpacas and llamas and provide a discussion of the important steps in the management of embryo transfer programs, expected results, and the factors affecting success rate.
Studies on reproductive biotechnologies (artificial insemination and embryo transfer) in alpacas and llamas were initiated in the early 1960s at La Raya Research Station, IVITA, San Marcos University, Puno, Peru. Novoa and Sumar (1968) reported the first surgical collection of embryos from the oviducts of alpacas following a fertile Alpaca donors may be sedated with Butorphanol tartrate (0.05–0.1 mg/kg IM) or a combination Xylazine (0.1 mg/kg IM) and Butorphanol (0.05 to 0.1 mg/kg IM). Alpacas are preferably placed in a sternal recumbency on an elevated platform. Epidural analgesia is provided by Lidocaine 0.2 mg/kg with a maximum of 1 ml per 50 kg of Body Weight) bifurcation of the uterine horns (Figure 28-1). The embryos collected 4 days after mating were at different stages of development from two cells to a morula. The recovery rate was 80% in single ovulating females. In another experiment using the same technique, Sumar and Franco (1974) obtained 44 morulae from donors superstimulated with equine chorionic gonadotropin (eCG; 700 international units [IU]) and induced to ovulate with human chorionic gonadotropin (hCG; 1000 IU).9 All embryos were transferred surgically to recipients, which resulted in a 10% pregnancy rate.
Figure 28-1 Illustration of the Surgical Embryo Collection Technique.
A, Syringe with blunt needle. B, Infundibulum. C, Oviduct. D, Curved forceps. E, Pipette inserted in the uterine horn. F, Glass dish. G, Uterine horn. (From Novoa, C., Sumar, J., 1968. Colección de huevos in vivo y ensayos de transferencia en alpacas. In: Tercer Boletín Extraordinario IVITA. Universidad Nacional Mayor de San Marcos, Lima, Perú, pp. 31–34.)
The first llama born via the nonsurgical collection and transfer technique was reported by Wiepz and Chapman (1985) in the United States.10 In this study, synchronization and ovulation of the recipient was induced by gonadotropin-releasing hormone (GnRH), and collection and transfer were done 7 days after GnRH treatment. One cria was born after transfer of two embryos to two synchronized females. Bourke et al. (1990) reported that superovulation can be achieved in llamas using pregnant mare’s serum gonadotropin (PMSG) and that embryos can be collected nonsurgically.11 From two llama donors, three embryos (all hatched blastocysts) were recovered by transcervical flushing 7 days after ovulation. Synchronization was achieved by simultaneous administration of 750 IU of hCG to both donor and recipients, immediately after mating of the donors. Two embryos were transferred to one of the recipients and one embryo to the second recipient. A single conceptus was visualized by ultrasonography in the recipient that had received two embryos.
Embryo transfer technology in alpacas and llamas has been relatively slow to develop until the last 10 years when renewed interest in the technology has led to more research on superovulation, embryo collection, and transfer.12,13 Commercial embryo transfer in alpacas and llamas is now offered in some countries, but it is still not fully accepted as a breeding technology in others.
Selection of donors should take into consideration the desired genetic traits as well as the absence of any possible genetic disorders. The genetic merit of the donor should be well above the average of the herd. Females should have good body conformation and health. No studies have been performed on the transmission of disease by embryos in llamas and alpacas, but it is important to screen all donors for common contagious diseases.
Although the technique may be considered in some cases of infertility, for maximum efficiency, donors should be reproductively sound. Breeding soundness evaluation of camelid females has been detailed elsewhere in this text. Ideally, adult females should have had at least one recent normal pregnancy and parturition.
The ideal recipient is a healthy parous female less than 8 years of age with normal reproductive function and lactation and mothering ability. However, maiden alpacas at 2 years of age and having reached at least 65% of adult body weight and height may be used as recipients. Recipients should have a body condition of 3 on a scale 1 to 5. Obese females and extremely thin females should not be used. All recipients should undergo a breeding soundness examination before selection for an embryo transfer program.
The recipient pool may include animals with poor fiber production and quality and those eliminated from breeding because of hereditary disorders that are not life threatening (blue eyes, multicolor coats, etc.). Lactating females may be used, but not earlier than 30 days after parturition. A recent experiment in the highlands of Peru on a large number of donors and recipients clearly demonstrated the negative effect of lactation and body condition on pregnancy rates.14 Pregnancy rates following embryo transfer were significantly higher in nonlactating recipient alpacas (44%) than in lactating ones (18.2%). Lactation was associated with poor body condition in this study, most likely because of negative energy balance and weight loss that may have interfered with corpus luteum function.
The availability of recipients is the most important limiting factor for the development of embryo transfer programs. In a program of fresh embryo transfer without superstimulation, at least three recipients should be available for each donor animal. In MOET programs, the best approach is to have a large pool of recipients to synchronize ovulation with the donor. New improved methods of embryo cryopreservation should help reduce the need for large numbers of recipients.
Donors should be mated when they present a 7- to 12-millimeter (mm) diameter follicle. We prefer to keep copulation duration to at least 8 minutes and not more than 20 minutes. GnRH (8.4 micrograms [mcg], intramuscularly [IM]) is administered immediately after mating. Serial transrectal ultrasound examinations are ideal for monitoring follicular development and making the decision to use the donor on the basis of follicular size. Occurrence of ovulation in the donor may be verified by measurement of serum progesterone on day 5 (day 0 = day of mating), ultrasonography, or teasing (nonreceptivity) for screening followed by ultrasonographic evaluation. Ovulation rate using these guidelines is usually greater than 80%.
In practice, alpaca and llama embryos are collected nonsurgically from the uterine cavity. The embryo recovery rate depends greatly on the timing of embryo descent into the uterine cavity following ovulation and fertilization. Detailed studies on the chronology of embryo development and descent into the uterus in alpacas and llamas are scarce. At 3 days after mating, embryos recovered surgically from the uterine tube were at various stages ranging from two cells to a morula.8 In alpacas, Bravo et al. (1996) using females that were slaughtered 4, 7, and 10 days after copulation reported that embryos were still in the uterine tube at the morula stage at 4 days. A compact morula and blastocysts were recovered from the uterus, at day 7 and day 10, respectively, after breeding.15
Cárdenas et al. (1997) reported the chronology of embryo development following mating in alpacas (day 2 = 2–4 cells embryos; day 3 = 4–8 cells; day 4 = early morula; day 5 = compact morula, early blastocyst; day 10 = collapsed or elongated blastocyst; day 11–15, elongated blastocysts).16 In this study, no embryos were recovered from day 6 to day 9, but the reasons were not known. More recently, Cervantes et al. (2008) reported that the alpaca embryo enters the uterus on day 6 after mating at the hatched blastocyst stage and grows rapidly to almost double the size between day 6 and day 7 after mating.17 In llamas, Taylor et al. (2000) reported recovery rates of 55%, 79%, and 100% at days 7, 8, and 10 after mating, respectively.7 Other studies in alpacas and llamas showed that hatched blastocyst can be collected from the uterine cavity at 6 to 6.5 days after mating with or without GnRH administration.11
A general agreement exists in the literature that alpaca and llama embryos reach the uterine cavity at the hatched blastocyst stage (Figure 28-2). The mechanisms controlling the descent of the embryo into the uterine cavity in camelids are still not understood. It is possible that hatching from the zona pelucida needs to occur within the uterotubal junction to permit passage of the embryo. The great variation in the reported chronology of embryo development and timing of descent into the uterine cavity may be attributed to the variability in the timing of ovulation and other factors influencing the speed of development.18 On the basis of our experience, the ideal time for embryo collection is 7.5 days after mating. This timing ensures the highest rate of embryo recovery (80% to 100%). Camelid embryos seem to develop more rapidly than those of other species and undergo elongation most likely as part of maternal recognition of pregnancy.18,19
Supplies needed for nonsurgical collection of embryos from alpacas and llamas are similar to those used in bovine embryo collection and consist of (1) a two-way Foley catheter (14 to 18 French [Fr]); (2) a stylette of 6.22 inch length with a clip to hold it within the catheter; (3) a sanitary plastic sleeve to prevent contamination of the catheter during introduction into the vagina; (4) a 6- or 12-mL syringe to inflate the Foley catheter balloon with the flushing medium; (5) an embryo filter placed on a graduated cylinder; (6) a “Y” tube for gravity irrigation of the uterus and return of the flushing medium (we suggest reducing the length of the tube, as commercial tubes are too large for uterine lavage in the alpaca or llama); and (7) searching and embryo washing dishes. All equipment should be sterile.
Embryo collection may be performed with the animal in the standing or sternal position in a chute. Sedation may be needed in some cases. In alpacas, embryo collection requires sedation and epidural analgesia. Alpaca donors may be sedated with Butorphanol tartrate (0.05–0.1 mg/kg IM) or a combination Xylazine (0.1mg/kg IM) and Butorphanol (0.05 to 0.1 mg/kg IM). Alpacas are preferably placed in a sternal recumbency on an elevated platform. Epidural analgesia is provided by Lidocaine 0.2mg/kg with a maximum of 1ml per 50kg of Body Weight). Another option is to use light sedation and physically restrain the donor in the chute.
The tail is wrapped and fixed dorsally to the fiber using clips of hemostat. Although most llamas may be easily palpated in this manner, provided ample lubrication is used, alpacas require more care and often only an operator with small hands may be able to accomplish this technique without harming the animal. After evacuating the rectum and locating the cervix and uterine horns, the hand is removed, the perineal area is scrubbed with water and soap, and the vulva is cleaned with disinfectant and dried.
The Foley catheter with its stylet is introduced aseptically into the vagina. Once the catheter is in the vagina, the operator places the hand into the rectum, locates the cervix, holds it gently between two fingers, and pushes it cranially, allowing the catheter closer to the cervical os. The catheter is gently threaded into the cervical canal until it is felt within the body of the uterus and is directed to the base of one horn. In alpacas, if direct transrectal palpation is not possible, the Foley catheter may be placed into the cervix with the aid of a vaginoscope. However, this technique is not always successful.
The Foley catheter is secured at the base of the horn by inflating the balloon (5 to 10 mL of flushing medium). The stylet is removed and a Y-junction tube is attached to the catheter, with one end attached to a bag of warm flushing medium and the return end connected to an embryo filter. The uterus is filled with the flushing medium until sufficiently distended (20 to 60 mL) and the return end is opened to recover the fluid into the filter. This procedure is repeated three or four times. It is important to allow the fluid to remain in the uterus for 30 to 60 seconds before recovering it. The procedure is repeated on the other horn. Alternatively, both uterine horns may be flushed at the same time by placing the balloon of the Foley catheter just cranial to the internal cervical os. A variety of flushing media that are commercially available for bovine and equine embryo transfer, for example, DPBS (Dulbecco’s phosphate-buffered saline) or Vigro (Bioniche, Pullman, WA) have been used in alpacas and llamas.
Embryo recovery rate for nonstimulated alpacas in the Mega Alpaca project was 74.2% (70.3% for the 2007 season and 78% for the 2008 season). These recovery rates were higher than in stimulated alpacas and llamas (50% based on number of corpora lutea).
Some potential problems which cause a low embryo rate recovery are the loss of the flush fluid when the balloon of the catheter is not fixed adequately at the end of the cervix, overmanipulation of the uterine horns, and the fluid entering the uterus with extreme turbulence. Soft latex gloves and generous amounts of lubricant or gel are recommended during transrectal maneuvering. When introducing the hand into the rectum, the operator must be very gentle and rotate his or her hand to pass the anal sphincter easily. The operator should never force the hand in. The ideal size of the operator’s hand is 20 centimeters (cm) or less in circumference, measured at the level of the carpal bones.
Donors are generally given a luteolytic dose of cloprostenol (250 mcg, IM) after flushing to prevent pregnancy in case of failure of embryo recovery. The rate of recovery of two embryos without stimulation varies from 2% to 6%.