Anatomy and Physiology of Reproduction in the Female Llama and Alpaca

Part 3: Reproduction CHAPTER

14  Anatomy and Physiology of Reproduction in the Female Llama and Alpaca Marcelo A. Aba

South American camelids (SACs) have for centuries been confined to the harsh and isolated environments of the Andes in South America. It is not surprising that little attention has been paid to them in comparison with the research conducted worldwide in other species. It was only during the last few decades that llamas and alpacas spread all over the world, gaining interest of the international scientific community because of their use as companion animals as well as for the production of high-quality fiber. This situation has drawn attention to the need for improvement in reproduction efficiency and the development of breeding techniques. Advances were mainly from the introduction of laparoscopy, the use of transrectal ultrasonography, and the availability of hormone assays. Nevertheless, current knowledge about the basic reproductive physiology of SACs is still incomplete. In their natural habitat, herd efficiency is considered to be very low because of reproductive failure. Attempts to introduce reproductive management programs and new reproductive biotechnologies at farm level (such as artificial insemination and embryo transfer) have been unsuccessful, in many cases because information from cattle and sheep was extrapolated to the species, neglecting their many unique reproductive features. Llamas, alpacas, guanacos, and vicuñas share many of their reproductive characteristics. This chapter presents an update on current knowledge in llama and alpaca female anatomy and reproductive physiology.

Female Reproductive Anatomy The ovaries of the llama are ellipsoid to globular (1.3–2.5 × 1.4–2.5 × 0.5–1.0 cm), while those of the alpaca are more globular (1.3–1.9 × 0.9–1.3 × 0.9–1.3 cm) varying the sizes according to the structures present on the ovaries (Figure 14-1).1 Each ovary weighs between 1.9 and 2.4 grams (g). In nulliparous females, the ovaries are flattened laterally and have an irregular surface because of the many small follicles present on it. In adults, numerous follicles, varying in size from 2 to 5 millimeters (mm), may be observed on the surface of the mature, normal ovary. They are not easily detected by transrectal palpation, especially in young animals, but are visually detectable by ultrasonography once they have reached a size of 3 mm.2 Larger follicles and corpora lutea that project from the surface of the ovaries can be easily detected by palpation and ultrasonography (Figure 14-2).3,4 Llama and alpaca cumulus oocyte complexes (COCs) are dark and easily

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distinguished through the follicle wall by trans-illumination. In llamas, the COC is located in the follicle hemisphere containing the expected ovulation point. Oocytes collected from follicles 2 to 11 mm in diameter range from 172 to 200 micrometers (µm). Immature oocytes had a very distinct and large germinal vesicle with a dark nucleolus. Mature oocytes display a metaphase plate surrounded by a dark area easily observable with low magnification.5 The ovarian bursa (a long conical, pocketlike fold of mesosalpinx) completely surrounds the ovary (Figure 14-3). Its apex forms a large circular orifice within which lie the fimbriae of the oviduct. The uterine tubes are rather long (10.5–18.3 and 20.4 ± 4.2 centimeters [cm] in llamas and alpacas, respectively), tortuous, quite firm, and embedded within the mesosalpinx. At the uterotubal junction, a papilla with a well-defined sphincter prevents the retrograde flow of fluid into the oviduct from the uterus, but the reverse is possible (Figure 14-4).1 The uterus of llamas and alpacas is bicornate, and in situ resembles the letter “Y” (Figure 14-5). The body is short (3–5.5 cm) and has approximately the same diameter. The left uterine horn may be slightly larger than the right one, even in nulliparous females. The length of the uterine horn of a llama, from the end of the septum to the tip of the horn on the convex surface, varies from 20 to 22.5 cm.1 The appearance of the uterine horns changes according to ovarian status. On ultrasonography, the uterus and the cervix appear heterogeneous because of an increase in intercellular fluid during follicular dominance. Meanwhile, cervical folds become more echogenic and prominent during luteal dominance. In general, the uterine horns are relatively straight and the uterine tone maximal during the estrogenic phase, whereas curling becomes maximal and the tone minimal during progesterone dominance.3,5 The uterus is located in the pelvic area in nongravid females, reaching the abdominal cavity during pregnancy. Considering that the majority of pregnancies are carried out in the left horn, asymmetry between both horns becomes extremely pronounced in multiparous females.4 The cervix is 2 to 5 cm long and 2 to 4 cm in diameter and has been described in two ways. One description indicates that two or three cervical rings are present (Figure 14-6). However, it has also been described as a single spiral fold with two or three turns, giving the appearance of two or three rings.1 The llama’s vagina, from the hymen to the cervix, varies in length from 15 to 25 cm and is approximately 5 cm in diameter.1 In alpacas, the vagina has an approximate length of 13.4

Chapter 14  •  Anatomy and Physiology of Reproduction in the Female Llama and Alpaca

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A

C B

Figure 14-1  Morphology of the alpaca ovary. Left pair of ovaries from

Figure 14-3  Ovary (B) exteriorized from its large bursa (A). Note the

a lactating anestrous female (Note old corpus albicans, arrow). Right set of ovaries from a cyclic female. Note the follicular wave on the same ovary. (Reprinted with permission, Tibary A and Anouassi A. Theriogenology in Camelidae, Second Edition, Actes Edition, in press, Actes Edition, Institut Agronomique et Vétérinaire Hassan II, Rabat Morocco.)

tortuous uterine tube (oviduct, C). (Reprinted with permission, Tibary A and Anouassi A. Theriogenology in Camelidae, Second Edition, Actes Edition, in press, Actes Edition, Institut Agronomique et Vétérinaire Hassan II, Rabat Morocco.)

Figure 14-2  Pair of ovaries from a female alpaca at 8 days after ovulation. Note the protruding large CL (15 mm) and the dominant follicle on the left ovary. (Reprinted with permission, Tibary A and Anouassi A. Theriogenology in Camelidae, Second Edition, Actes Edition, in press, Actes Edition, Institut Agronomique et Vétérinaire Hassan II, Rabat Morocco.)

± 2.0 cm and a diameter of 3.4 ± 0.7 cm. The vulva is small, with an opening of 2.5 to 3 cm in llamas, and has well-defined external labia that lie in a slightly slanted to vertical position approximately 4 to 6 cm ventral to the anal orifice.1 The vulva remains virtually unaltered despite changes in ovarian structure or sexual receptivity.

Female Reproductive Physiology Puberty The precise age at onset of puberty in female llamas and alpacas remains undetermined. Age at first ovulation depends on age at first mating as camelids are induced ovulators.6 Current knowledge suggests that puberty is greatly influenced by nutritional status and environmental conditions. In their

Figure 14-4  Open left uterine horn. Note the protruding uterotubal junction papilla (arrow) at the tip of the horn. (Reprinted with permission, Tibary A and Anouassi A. Theriogenology in Camelidae, Second Edition, Actes Edition, in press, Actes Edition, Institut Agronomique et Vétérinaire Hassan II, Rabat Morocco.)

natural habitat, the puna, age at puberty is reflected by forage availability and time of birth relative to the wet season. Nutritional deprivation during early development delays puberty. An increased production of ovarian hormones has been observed at approximately 6 months of age in well-fed llamas. Ovarian activity has been reported to begin around 10 months of age in alpacas, by which time the diameter of follicles is 5 mm or larger. In addition, it has been demonstrated that the age at which breeding maturity is reached depends partly on body weight. The first successful mating in female alpacas may occur at 12 to14 months of age and a body weight of approximately 40 kilograms (kg). Furthermore, a relationship seems to exist between body weight at mating and ability to maintain pregnancy. For each kilogram of gain in body weight, a 5% increase in parturition rate is seen. However, pregnancy rate becomes independent of body weight beyond 33 kg. Under conditions of good nutrition, a body weight of 33 kg is easily

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A B

C

achievable in most yearlings. The usual practice in the puna is to breed llamas and alpacas for the first time at 2 years of age, although pregnancies have been observed in 6-month-old females. In any case, it is recommended that breeding be delayed until the female has reached two thirds of her anticipated adult body weight to reduce risks of stunted growth of the dam and dystocia.4

D E F

G H

I J

Figure 14-5  Anatomy of the reproductive tract in a female alpaca. uterine horn (a), note the absence of clear bifurcation externally; left ovary (b), brad ligament (c), fornix and external os of the cervix (d), urinary bladder (e), vagina (f), vestibule-vaginal sphincter (g), urethral orifice (h), vulvar lip (i), clitoris (j). (Reprinted with permission, Tibary A and Anouassi A. Theriogenology in Camelidae, Second Edition, Actes Edition, in press, Actes Edition, Institut Agronomique et Vétérinaire Hassan II, Rabat Morocco.)

B

A

Figure 14-6  A, Cervix; note the three cervical spiraling rings. B, Body of the uterus is very small because of the long septum separating the uterine horns. (Reprinted with permission, Tibary A and Anouassi A. Theriogenology in Camelidae, Second Edition, Actes Edition, in press, Actes Edition, Institut Agronomique et Vétérinaire Hassan II, Rabat Morocco.)

Seasonality Whether SACs are seasonal breeders or not is still being debated. This seems to be related to environmental constraints and management systems. In their natural habitat, the harsh highlands of the Andes, they usually display sexual activity during the warm and rainy months from November to April, when food availability increases naturally. Consequently, parturitions take place also during the next rainy season, which, in turn, implies that breeding occurs in the same period again. Conversely, when animals are reared in milder habitats, and nutritional requirements are fulfilled, llamas and alpacas are sexually active throughout the year, and females may give birth at any time of the year. SACs are therefore classified as nonseasonal breeders, although environmental or nutritional factors may force them to behave as seasonal breeders.

Ovarian Activity Llama and alpaca females are induced ovulators, requiring copulation to trigger the luteinizing hormone (LH) surge responsible for ovulation.7 Therefore, they do not have regular estrous cycles, as described in other domestic species. Instead, it is advisable to refer to ovarian activity in terms of follicular phase and luteal phase. Studies using laparoscopy, ultrasonography, and hormone profiles in llamas and alpacas have demonstrated the occurrence of repetitive cycles (waves) of ovarian follicular growth and regression without ovulation in absence of copulation.8,9 In fertile, nonmated females, during each follicular wave, a cohort of follicles is recruited, followed by the synchronous emergence of a group of antral follicles, which grow to 4 to 5 mm in diameter. Of these, one follicle is selected to become the dominant follicle, and the others undergo atresia and regress.3 The dominant follicle grows to maturity and finally regresses if not ovulated. The dominant follicle exerts a negative effect on the rest of the follicles in both ovaries, thus regulating their number and diameter. Proposed mechanisms involved in the suppression of follicular growth by dominant follicles include reduction in pituitary gonadotrophin (follicle-stimulating hormone [FSH]) release because of high estradiol plasma concentrations, locally produced substances such as inhibin and androgens, or both, as it has been described in other species. Before final regression of the dominant follicle, a new follicle emerges and starts growing, producing the coexistence of regressing and growing follicles over several days, a phenomenon known as overlapping follicular waves. In most nonovulating llamas, waves overlap about 30% of their lifespan, that is, two successive follicular waves coexist for approximately 8 days. Because of this peculiarity, a follicle larger than 5 to 6 mm in diameter may be found at any given time in camelids, although it is not possible to discern whether the observed follicle is growing or regressing at a single examination.

Chapter 14  •  Anatomy and Physiology of Reproduction in the Female Llama and Alpaca In nonlactating llamas, the duration of follicular waves varies between 20 and 25 days.9,10 The follicular growth phase spans, on average, 9 days; the plateau phase takes approximately 5 days; and the regression phase lasts 8 days. The interwave interval is defined as the time elapsed between the beginnings of two consecutive waves. This period ranges from 16 to 20 days in llamas. However, because of the high variation observed in this interval in both species, it has been recommended that use of a mean value be avoided, as it does not accurately describe what is occurring in an individual animal, nor does it allow prediction of the optimal time for breeding.6 During the growth phase, follicles grow at rates of 0.5 to 0.9 mm/day.9 Ovulatory size is attained 6 to 8 days after the beginning of the growth phase. The maximum diameter of the dominant follicle averages 12 mm (range 9–16.5 mm), and this size is reached approximately 12 days after the beginning of growth. Thereafter, follicle regression proceeds at a rate of 0.8 mm/day. Lactation and pregnancy strongly affect follicular activity, and their negative effects are additive in llamas, conditioning shortened interwave intervals and reduced maximum diameter of dominant follicles. Thus, during gestation, waves begin at 15-day intervals, and dominant follicles reach a maximum diameter of 10 mm.10 In alpacas, maximum follicular diameters average 9 mm (range from 8 to 12 mm). On the basis of laparoscopic examinations of the ovaries, the average duration of follicle development, maintenance, and regression was consistently estimated at approximately 12 days.8 More recent studies carried out in Australia have reported interwave intervals ranging from 12 to 22 days.11 This indicates that the duration of follicular waves is widely variable (probably ranging from 12 to more than 20 days). Despite the high variation reported, the growth rate of dominant follicles seems to be consistent over the first 10 days after emergence in alpacas averaging 0.4 mm/day.11 In addition, longer interwave intervals were associated with the development of larger-sized follicles. It remains to be determined whether phylogenetic links exist between alpacas and vicuñas, a species in which the interwave interval averages 4.2 days for nonpregnant, nonlactating animals and if possible influence of crossbreeding with llamas explains this individual variation among alpacas.12 As in llamas, the mature phase lasts between 2 and 8 days, and the regression phase takes 3 to 8 days.9,11 In both species, the occurrence of large dominant follicles between ovaries is almost equal.11,13 No regular pattern of dominant follicles emerging between the left ovary and the right ovary has been demonstrated. The 65% alternation between ovaries reported in one study indicates that many successive waves occur in the same ovary.9 The view of the two ovaries as one unit controlled by systemic rather than local mechanisms has been proposed. A close relationship between follicle size and estradiol-17β secretion has been reported.9 In addition, peak estradiol-17β plasma concentration occurs approximately 12 days after the beginning of the growing phase in llamas and 8 days in alpacas and is closely associated with the maximum size of the follicle.4 Plasma progesterone concentrations remain basal in anovulatory animals, offering an interesting animal model to study follicular dynamics in the absence of progesterone. Given that follicular waves usually overlap, the estrogen produced during those waves remains elevated, determining long

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periods of receptivity (≥30 days) interrupted by short stages of male rejection.8,10 In both species, follicles are considered ovulatory, and copulation occurs over a window of several days. Available evidence indicates that growing follicles attain ovulatory capacity when they reach a diameter of 7 mm. Some authors have suggested that smaller follicles may ovulate, but these results are still debated. Follicles may be induced to ovulate during the rest of the growing phase, the plateau phase and beginning of the regressing state. However, large regressing follicles may undergo luteinization rather than ovulation. Although male rejection is usually indicative of low plasma estrogen concentrations, male acceptance is not associated with the presence of an ovulatory follicle in camelids. Therefore, many matings performed on the basis of receptivity may be infertile because of lack of ovulation. In addition, the fact that ovulation might be induced in camelids from a wide range of follicle sizes and the subsequent maturational status raises the question about the effect on quality of the ovum and its effect on embryo survival. The mechanism controlling growth and recruitment of follicles in SACs has not yet been elucidated. Studies in sheep indicate that follicle recruitment requires at least basal levels of gonadotropins. In cattle, the secondary surge of FSH following ovulation may play a role in initiating follicular recruitment in the next cycle, and a relationship has been proposed between FSH and the emergence of follicular waves. Thereafter, estrogens and inhibin secreted by a developing follicle inhibit further central FSH secretion, which becomes inadequate for the growth of subordinate follicles but stimulates growth and cell differentiation within the same follicle. No temporal association between FSH secretion and follicle development has been observed in llamas.13 Further studies, including development of sensitive methods for FSH measurement, are needed for better understanding of the process of follicular recruitment in llamas and alpacas. Anovulatory follicles characterized by a larger diameter and a longer lifespan have been described in llamas and alpacas.14 Structures exceeding 12 or 14 mm in alpacas and llamas, respectively, and lasting 25 to 30 days have been referred to as cystic follicles. Follicular development appears to be normal through initial growth and maturity, with the follicle continuing to grow over the expected maximum diameter without any transition period between maturity and cystic status. In studies where the gross and ultrasonic morphologies of persistent follicles were evaluated, similar structures were identified as hemorrhagic follicles. They appeared grossly as a large fluctuating structure, filled with a dark red content, and ultrasonically they showed free floating echogenic content.15 Occurrence of anovulatory, oversized follicles appears increased in nonmated animals, and some individuals seem to have a higher tendency to develop them. Whether cystic and hemorrhagic follicles are the same structures or not is still debated. These structures do not seem to secrete estrogen. They are refractory to, and their presence is not associated with, any alteration or abnormality of ovarian dynamics.

Copulation and Mating Behavior Behavior related to the establishment of dominance hierarchies is observed among males when they are first placed

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together with females. This behavior consists mainly of challenging and is characterized by spitting and jumping. Dominance is usually established quickly. The initiation of breeding behavior is marked by the pursuit of females by males. Some females become recumbent immediately, but most move away from the male until he mounts and puts pressure on the hindquarters in an attempt to force the female to lie down in a seated position. It is important to point out that sitting position is also assumed by females in response to the stress of being chased by the male. Once the female is recumbent, the male mounts her and moves forward to achieve intromission. Males penetrate the cervix during copulation and deposit semen into both uterine horns.16 During copulation, males show their excitement with trembling ears, tail flipping up and down, nostrils dilating, and constantly vocalizing by making a guttural sound. Females assume a very passive attitude and, in some instances, may change position and lie in lateral recumbency. Copulation generally lasts from 3 to 65 minutes, with an average time of 20 to 30 minutes.17 Ejaculation is a continuous process during mating. Receptive females generally sit near a mating couple. Nonestrous females strongly reject the males by spitting, kicking, and running away, but some may assume a sternal posture and submit to mating by a dominant male.1 Sexual activity is particularly intense during the first week following the introduction of males into a herd of nonpregnant females. More than 70% of the females are mated at least once during this time. Thereafter, the continuous association of males and females inhibits the sexual activity of the males in spite of the presence of estrous females. Although the physiologic basis for the decline remains unknown, it has been shown that if the males are allowed to rest for 4 to 5 days, they resume normal sexual activity. By taking such simple measures, conception rates have been increased from the usual 50% to 70%.18 The presence of a functional corpus luteum (CL) is usually associated with a nonreceptive status. During the luteal phase, the female strongly rejects the male by spitting, kicking, screaming, and attempting to escape.

Ovulation Ovulation is defined as the process by which a mature ovarian follicle ruptures and releases an ovum (oocyte). In camelids, usually only one oocyte is shed at each ovulation from any point at the surface of the ovary except from the hilus. Pulsatile release of gonadotropin-releasing hormone (GnRH) from the hypothalamus and the consequent LH release from the hypophysis are involved in the ovulatory process in mammals. Species are usually classified as spontaneous or induced ovulators, depending on the stimulus involved in triggering the process. In most domestic species, the growing follicle in each cycle releases increasing amounts of estradiol, which, is responsible for eliciting a GnRH surge once a threshold concentration is exceeded. This ensures that ovulation occurs in temporal association with signs of heat and male acceptance. Conversely, in induced ovulators such as camelids, the hypothalamus is not responsive to estradiol concentrations, requiring neural signals associated to mating activity to elicit the GnRH (and subsequently LH) peak responsible of ovulation. A classic study evaluating the effect of different mating stimuli on ovulation induction in alpacas showed that animals mated by intact or vasectomized males ovulated in higher proportion

(80%) compared with those only mounted (15%), indicating that mounting and penile intromission are both necessary to provide adequate stimulus for LH release and subsequent ovulation. Conversely, no relationship has been observed between duration of copulation and ovulatory response, as repeated mating does not improve ovulation rates. A 100% ovulatory response has been reported after a single injection of human chorionic gonadotrophin (hCG) in alpacas.19 GnRH and LH injections have proven to be equally effective in both species. Evidence has indicated that a factor present in the semen is involved in the induction of ovulation in the Bactrian camel, since ovulation occurs after intravaginal or intramuscular or intrauterine administration of seminal plasma from the species.20,21 Similar observations indicate the existence of an ovulation-inducing factor (OIF) in the seminal plasma of llamas and alpacas. Alpaca seminal plasma stimulates LH secretion from primary cultures of rat pituitary cells, which suggests the presence of a factor or factors in alpaca seminal plasma that had GnRH-like activity.22 This factor appears able to elicit a surge in LH plasma concentrations and induce ovulation in more than 90% of treated animals when injected intramuscularly; this gives further support to the hypothesis of a systemic rather than a local mechanism.23 Whether intrauterine infusion is effective at inducing ovulation is still being debated. Although some studies failed to induce ovulation using the intrauterine route consistently with the observation that ovulation rate is not increased by artificial insemination, others have reported that ovulation was registered in 6 of 10 alpacas and 5 of 8 llamas after intravaginal deposition of alpaca semen.23 However, in three separate studies in Bactrian camels, ovulation was induced by intravaginal or intrauterine infusion of whole semen or seminal plasma in 75% of females. This difference might partially be related to the fact that a normal consequence of copulation is an inflammation of the endometrium as a result of repeated abrasion by the penis. Hyperemia of the excoriated endometrium might facilitate absorption of the factor, as was evidenced in a study in which disruption of the endometrial mucosa by curettage increased the ovulatory effect of seminal plasma (67% of females ovulated compared with 43% in the group that received intrauterine infusion and 93% in the intramuscularly treated animals). Attempts to identify the nature of the molecule in llamas indicated that the molecule is a protein whose molecular weight is roughly greater than 30 kilodalton (kDa).24 This is consistent with a report on Bactrian camels in which the molecule was suggested to be a folded complex of greater than 50 kDa. The OIF has been identified recently as β-nerve growth factor.25,26 The first increase in LH concentrations in peripheral blood occurs about 15 minutes after the initiation of copulation in SACs, and peak concentrations (around 10 microgram (mcg)/ L−1) are registered 2 to 3 hours after copulation. Plasma LH concentrations remain high 6 hours after copulation and drop to basal concentrations by 12 hours after mating.27 A similar response has been observed after treatment with GnRH.28 In addition, repeated copulation or GnRH injection does not induce further release of LH. Interestingly, the LH secretory pattern observed after seminal plasma injection appears delayed compared with that registered in GnRH-treated animals. The first significant increase occurs 1 hour after injection of seminal plasma, and the maximum concentration is reached at 2 hours, with the first significant decrease occurring 2.5 hours later than in the

Chapter 14  •  Anatomy and Physiology of Reproduction in the Female Llama and Alpaca GnRH group.23 These observations give further support to the hypothesis that OIF is biochemically different from GnRH. This is consistent with the observation that the LH-secreting effect of alpaca seminal plasma on rat pituitary cells in vitro is not suppressed when anti-GnRH antibodies are added to the primary culture.22 In parallel, a twofold rise in the mean prostaglandin F2α (PGF2α) metabolite plasma concentration has been recorded immediately after copulation (from 40 ± 4 picomole per liter [pmol/L−1] before copulation to 81.3 ± 14.6 pmol/L−1 within 5 minutes after copulation). Thereafter, PGF2α metabolite plasma concentrations decline rapidly and reach values similar to those recorded before copulation by 4 hours after mating. Similarly, increased cortisol plasma concentrations are registered in all animals after copulation, reaching peak concentrations around 2 hours after mating and remaining high for at least 12 hours after copulation.27 It could be speculated that the prostaglandin secretion might be related to a potent Fergusson reflex, involving oxytocin secretion from the neurohypophysis. The cortisol release associated with copulation might be related to the stress of mating behavior (pursuing for several minutes, long mating times, etc.). However, any relationship between the secretory mechanisms of these hormones and their possible roles in the ovulatory mechanism of llamas remains to be elucidated. The LH surge triggers resumption of meiosis in the oocyte, disruption of cumulus cell cohesiveness, and rupture of the follicular wall.6 Elevated estradiol-17β plasma concentrations are registered during the day of mating and remain elevated 24 hours later, which explains the frequent observation of females accepting the male the day after first mating. Estradiol concentrations drop to basal values within 2 to 3 days after copulation, in association with the disappearance of the dominant follicle because of ovulation.29,30 No significant differences are observed in the preovulatory LH peaks (either in amplitude or in duration) and in the secretory profiles of progesterone, estradiol-17β, cortisol, and PGF2α metabolite in response to copulation with an intact or a vasectomized male or between matings with an intact male, whether or not pregnancy resulted.27 Ovulation has been reported to occur approximately 30 hours (range 24–48 hours) after copulation in SACs.31 Similar intervals between treatment and ovulation have been reported in animals treated with LH or GnRH.31 Induction of ovulation and quality of the ensuing CL are affected by the size of the follicle at mating. Bravo et al. stated that copulation only results in ovulation and development of a normal CL from mature follicles between 7 and 12 mm in size.29 Animals with small growing follicles seem to release approximately half the amount of LH over a 6-hour postcopulatory period compared with those with large follicles, and subsequent follicular development is not interrupted in response to copulation. In animals with regressing follicles at the time of mating, luteinization—not ovulation—occurs, although copulationinduced LH release is similar to that in animals with mature follicles.29 Soon after ovulation, a new follicular wave starts. Progesterone from the CL exerts a negative influence on growing follicles whose final maturation occurs after luteolysis. The frequency of ovulation failure after natural mating has been reported to be between 10% and 30%.32 Multiple ovulations occur in 8% to 10% of the animals.4 Although, it is

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widely accepted that camelids are induced ovulators, spontaneous ovulations have been observed in both llamas and alpacas, with an incidence of 3.5% to 10%.8,33 The mechanisms involved in spontaneous ovulation have yet to be studied.

Corpus Luteum Formation and Function After ovulation, the ruptured follicle undergoes a dramatic transformation into the corpus luteum, which is a steroidogenic cluster of cells responsible for the secretion of large amounts of progesterone. The newly formed CL can be clearly detected with laparoscopy 3 to 4 days after mating in alpacas and with ultrasonography 4 to 5 days after mating (2–3 days after ovulation) in llamas. In a study on 928 alpacas, 50.9% of corpora lutea were located in the right ovary and 47.4% were in the left, whereas 1.7% of the animals displayed a functional CL on each ovary.34 The CL reaches a maximum size (12–14 mm) on day 8 or day 9 after mating in nonpregnant animals and declines sharply thereafter. Complete regression of the CL has been reported between days 15 and 18.15 In pregnant animals, maximum diameter (approximately 16 mm) is attained on day 21.4 ± 1.2. It has been shown that the CL is necessary for the maintenance of pregnancy throughout the gestation period in both species.35 A close temporal relationship exists between the morphologic changes of the CL detected by ultrasonography and the changes in plasma progesterone concentrations, except during luteolysis when progesterone decreases 1 to 3 days before the morphologic changes can be detected.15 The increase in plasma progesterone concentrations above basal levels is detected on day 4 after mating and reaches peak concentrations on day 8 or day 9 after copulation in llamas. A similar progesterone pattern is observed in alpacas; however, the increase by day 4 is more pronounced in alpacas than in llamas. Plasma progesterone concentration drops sharply to levels recorded before ovulation by day 10 or day 11 because of the release of PGF2α from the uterus. Considering the period of more than 1 day from mating to ovulation, the CL lifespan is estimated to be 9 days in camelids.36 Females should be sexually receptive again approximately 12 to 14 days after mating if conception does not occur. Pseudopregnancy, a common feature in induced ovulators after sterile mating, is rarely observed in llamas and alpacas. The developmental kinetics of the CL induced by LH or GnRH injection are similar to those induced by natural mating in terms of CL diameter, plasma progesterone concentrations, and onset of regression. Thus, hormonal preparations (LH and GnRH) appear equally reliable for inducing ovulation and normal luteal function, and both are suitable for use in synchronization for artificial insemination protocols or embryo transfer programs.31 Recent studies have shown that CL diameter and progesterone levels are higher after induction of ovulation by seminal plasma (OIF) than by GnRH injection.

Luteolysis Luteolytic pulses of PGF2α start on day 7 or day 8 after mating (approximately day 5 or day 6 after ovulation). Luteolysis is completed by day 9 or day 10 after mating in nonpregnant llamas and alpacas, although prostaglandin pulses continue until day 13. The first peaks, registered on days 7 and 8 have

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LH

PGF2

Estradiol

Progesterone

Mating

Luteolysis

New follicular phase

Figure 14-7  Schematic drawing of endocrinologic events following a sterile mating. Estradiol decreases after mating. Luteinizing hormone peaks a few hours after mating and induces ovulation. The corpus luteum reaches its maturity at 8 days. Luteolysis occurs 9 to 10 days after mating and a new follicular wave (receptivity resumes 12 to 14 days after the sterile mating.) consistently lower amplitudes than those recorded during the following days. It is worth mentioning that the camelid embryo reaches the uterus by day 6.5 or day 7, which coincides with the first PGF2α pulses. Calculation of the area under the peaks shows that about 35% of the total release of PGF2α occurred during days 8 and 9, when luteolysis was at its highest, whereas about 65% of the total release is seen after progesterone levels have decreased below 1 nmol/L−1. The transfer of PGF2α, from the utero-ovarian vein to the ovarian artery seems to involve a countercurrent mechanism similar to that observed in other ruminants. With regard to the molecular events driving the luteolytic process, exposure to an environment characterized by high progesterone concentrations is claimed to play a key role in the regulation of uterine oxytocin receptors and, consequently, of prostaglandin pulsatile release in other species. Considering that in SACs, the first significant increase in progesterone concentrations is observed 3 to 4 days after mating, such a period lasts only 3 to 4 days in llamas, whereas it lasts around 3 and 5 times longer in sheep and cows, respectively (Figure 14-7).30 Recent studies have demonstrated that maximal expression of estrogen and progesterone receptors is attained during the follicular phase. A drop in estrogen receptor–α expression is observed after ovulation (day 4 after induction of ovulation), increasing again at the end of the luteal phase when plasma estrogen concentrations start to increase because of the beginning of a new follicular growth. After ovulation, progesterone receptors expression steadily decreases, being minimal when luteolysis occur. By that time, in relation to the release of luteolytic pulses of PGF2α, a sharp increase in the expression of cyclo-oxygenase 2 (the limiting enzyme in the synthesis of PGF2α) is recorded. No relationship between the expression of estrogen receptor–α and oxytocin receptor have been detected. Oxytocin receptor expression remains at the same level during the follicular and luteal phase. No significant differences in endometrial receptors have been observed between both uterine horns.37,38 Even though ovulation frequency is nearly equal between both the ovaries, almost all successful pregnancies must be

established in the left uterine horn.34 A difference in luteolytic effect between the right (local) and left (both local and systemic) uterine horns has been proposed as the reason for this unique trait. It has been demonstrated that removal of the left horn prolongs luteal activity in animals carrying a CL in the left ovary. Conversely, removal of the right uterine horn in those females with a functional CL in the right ovary causes only a slight delay in luteal regression. Thus, an embryo derived from an ovulation in the right ovary must migrate into the left uterine horn; otherwise, the empty left uterine horn will cause luteolysis in the right ovary and terminate the pregnancy.39 The particular angioarchitecture of the reproductive tract in SACs has been proposed to be responsible for this uniqueness. Unlike in other mammals, a major branch of the right uterine artery crosses the cranial intercornual area to supply much of the left uterine horn, whereas a corresponding major vein originates from the left horn, crosses the midline, and terminates as a branch of the right uterine vein. The presence of a large cross-over venous drainage from the left horn to the right side is compatible with the hypothesis that the left horn can exert luteolytic control over the CL in the right ovary through a venoarterial pathway.40 However, a similar study carried out in dromedary camels (a species also carrying almost 100% fetuses in the left horn), failed to detect such cross-over system, which suggests that some other component might be involved in selecting the left horn for implantation.41 In addition, the question that remains is why the pregnancy with a CL in the right ovary is not terminated by the local effect of prostaglandin released from the empty right horn. It has been shown that luteal lifespan in camels can be artificially prolonged if they are treated daily with meclofenamic acid, a prostaglandin synthetase inhibitor, starting on day 6 after ovulation.42 A similar study conducted in llamas showed that meclofenamic acid can extend the luteal phase in approximately one third of treated animals.37 A similar approach using flunixin meglumine proved to be effective in depressing prostaglandin synthesis in llamas. This negative effect was not strong enough to completely inhibit the occurrence of the luteolytic pulses of PGF2α, and luteolysis proceeded in these animals. However, based on progesterone patterns, luteolysis seems to be delayed by 1 to 1.5 days in flunixin meglumine-treated llamas.30 In llamas and alpacas, although progesterone concentration from ovarian origin remains elevated throughout pregnancy, a transient decrease and subsequent recovery in progesterone are observed after day 9 after mating. In connection with the progesterone decline, PGF2α pulsatile release has been observed in all pregnant animals from day 7 to day 15 after mating. During this period, prostaglandin peaks are recorded in all pregnant llamas at the time when maternal recognition of pregnancy (MRP) is expected to occur. The magnitude and the frequency of the pulses registered between days 7 and 12 of pregnancy are not comparable with those observed in nonpregnant animals, averaging about 3% of the total amount of prostaglandin released during luteolysis. Nevertheless, this prostaglandin production proved to be effective in inducing a decline in progesterone concentrations. A similar transient decrease in progesterone concentrations associated to PGF2α pulsatile release during MRP is observed in pregnant alpacas.30,43 In addition, the endometrial expression of cyclooxygenase 2 in luminal epithelium decreases during MRP and

Chapter 14  •  Anatomy and Physiology of Reproduction in the Female Llama and Alpaca by day 12 after mating is a third of that reported during luteolysis in nonpregnant animals.37

Maternal Recognition of Pregnancy Available information indicates that the MRP occurs earlier (days 8 to 10 after mating) in SACs than in ruminants and encompasses a very short period. The process of MRP remains poorly understood in camelids. On day 11 after estrus in sheep and day 13 in cows, the blastocyst undergoes a logarithmic elongation phase, a process intimately related to the MRP occurring on days 12 and 16, respectively. In camelids, the allantois elongates after migration and soon protrudes from the left horn into the uterine body and the right horn.44 The resurgence of progesterone and rescue of the CL suggest that a signal produced by the blastocyst in this period of gestation is responsible for the MRP. The mechanism by which the pulsatile release of PGF2α from the uterus is depressed (but not suppressed) in the pregnant llama and alpacas is not well understood. Attempts to identify any substance similar to ovine or bovine interferon-τ (IF-τ) in camel conceptus incubates from days 10 to 33 after ovulation, have been unsuccessful.45 No increase in estrone sulfate concentrations in urine or estradiol-17β in plasma has been recorded during early pregnancy in SACs.36,46 However, preimplantation llama blastocysts produce increasing amounts of estradiol-17β during days 7 through 15 of pregnancy.47 Similarly, a considerable aromatizing ability of the extraembryonic membranes as early as 10 days after ovulation has been reported in camels.48 On day 7, llama blastocysts are spherical (90%) or ovoid (10%) in shape. The dramatic increase in blastocyst estradiol production observed between days 11 and 13 occurs when they are transforming from an ovoid morphology to an elongated morphology. This increase in estradiol production by the llama blastocyst occurs in temporal association with the period proposed for MRP. Moreover, estradiol administered to ovulated females transiently prolonged luteal lifespan and increased progesterone secretion during the period when luteolysis would have been completed.47 In addition, 200 mg of estradiol injected on days 8 and 9 after ovulation increased embryo survival from 57.1% to 86.7 % in control and treated animals respectively, suggesting its possible involvement in signaling MRP in camelids.49 A recent study has demonstrated an increase in the population of estrogen receptors–α between days 8 to 12 after mating in pregnant animals, giving further support to the hypothesis that estrogens from the embryo are involved in the process of MRP in camelids. In addition, a reduction in the expression of progesterone receptors has been reported by day 12 after mating.37 It has been suggested that in other species, the loss of progesterone receptors during early pregnancy is a key factor to allow the expression of proteins necessary for nutrition and implantation of the embryo. In conclusion, MRP appears to be an extremely complex and tightly timed process in camelids, and the chances of embryo loss around this period increase significantly.

Gestation The normal length of gestation ranges from 335 to 360 days in llamas and averages 327 and 346 days for Huacaya and

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Suri type alpacas, respectively.50 Twining is extremely rare in camelids.

Early Embryo Development The events leading to fertilization in camelids remain poorly understood, although it is assumed they might be similar to the processes studied in other domestic animals. Thus, the oviduct is supposed to play a key role in sperm motility, viability, and fertilizing capacity. Considering that ovulation occurs several hours after mating in camelids, it is highly probable that a sperm reservoir is maintained in the oviduct while the release of the egg is awaited. During storage, the fertile lifespan of spermatozoa is maintained by binding to the epithelial cells lining the oviductal lumen. Current evidence strongly suggests that the uterotubal junction is a good candidate for acting as a sperm reservoir. Its anatomic features and the presence of a bonding agent in this region support the assumption.51 After fertilization, embryos develop rapidly in llamas and alpacas. Four days after mating, a 4- to 8-cell morula may be found in the oviduct.52 Embryos enter the uterine cavity approximately 6 to 7 days after mating (5 to 6 days after ovulation) at the hatching or early hatched blastocyst stage.5 At this stage, embryos are spherical, exhibiting great variation in the diameter. Embryo size ranges from 450 µm or less to 651 µm or greater in diameter in llamas and alpacas. Thereafter, trophoblast expansion ranges from a mean of 1.2 mm in diameter on day 6.5 to 7.5, beginning of elongation by days 9 to 10, to 83 mm in length on day 13 to 14.2 The trophoblast establishes close contact with the endometrium by day 12 of pregnancy (Figure 14-8).53 This accelerated rate of embryo development may be related to the apparent early MRP in these species.

Endocrinology of Gestation The CL is necessary for the maintenance of pregnancy throughout the gestation period in llamas and alpacas.35 After its transient drop, around the expected time of MRP, plasma progesterone concentrations remain elevated throughout pregnancy, begin to decline between 15 days and 1 day before parturition and finally drop from day 1 before parturition until parturition.54 Increasing prostaglandin concentrations are observed from mating onward, and peak concentrations are attained between weeks 7 and 10 of pregnancy in llamas and around week 8 of pregnancy in alpacas. During the last 90 days of pregnancy, plasma concentrations of 15-ketodihydroprostaglandin F2α metabolite increase steadily until the day of parturition. In both species, plasma estrone sulphate concentrations remained below 1 nmol/L−1 throughout the first 8 months of pregnancy, after which time they rose steadily, with the highest values recorded during the last 20 days of pregnancy. Thus, in llamas and alpacas, as in other species, it is possible to confirm that pregnancy is in progress by estrone sulphate analysis during the last 90 days before parturition. Plasma estradiol-17β concentration remains elevated during the last 45 days of pregnancy. The first significant decrease in estradiol concentrations is observed during the day of parturition.

Placenta A complete account of the structure of the placenta in camelids has been given by Olivera et al. and will not be presented in depth here.55 In brief, the placentation of the camelid has

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Part 3  •  Reproduction

B

A

C Figure 14-8  A, Blastocyst 5 days. B, Hatched blastocyst. C, Elongated embryos 12 days. (Reprinted with permission, Tibary A and A Anouassi A, Theriogenology in Camelidae, second edition, Actes Editions, Institut Agronomique et Veterinarie Hassan II, Rabat, Morocco.)

been classified as diffuse, epitheliochorial, and microcotyledonary, similar to that of the mare and the sow, with some uniqueness. In late gestation, the intercapillary distance across the diffusion pathway has been reported to be as narrow as 2 µm. This is assumed to be one of several adaptations to carrying out pregnancy at high altitude. Before day 15 of pregnancy, the elongated blastocyst remains completely free within the uterine lumen. Implantation starts sometime between days 15 and 22 of pregnancy. On day 22, a maternal–embryonic interaction becomes macroscopically evident. By that time, complete interdigitations between the membranes of uterine epithelial and trophoblastic cells are observed, especially in the left horn in areas around the embryo, whereas a weak apical contact is present in the rest of the left horn and in the right horn, where implantation proceeds later on. The process seems to extend from the left to the right horn following a wavelike pattern. By day 45, the depth and complexity of interdigitations have increased significantly, and the embryo becomes firmly attached to the maternal tissues. In parallel, secretory granules are observed in the uterine epithelium, and giant multinucleate cells appear interposed between the remaining trophoblast cells, showing intense alkaline phosphatase activity, deposits containing iron and periodic acid-Schiff (PAS)–positive granules. Attempts to detect placental lactogen hormone have been unsuccessful both in binucleate and multinucleate trophoblast cells. By day 45 of gestation, the extraembryonic connective tissue becomes well vascularized, indicating the beginning of placentation. Current data suggest that although the

epitheliochorial alpaca placenta is diffuse, various trophoblast cell types and specialized areas of the maternal–fetal interface give the placenta micro-regional functions, in which histiotrophic nutrition, hormone production, and molecular exchange are prevalent.55 In late gestation, the intercapillary distance across the diffusion pathway has been reported to be as narrow as 2 µm. This is assumed to be one of several adaptations to carrying out pregnancy at high altitude. Another peculiarity of the SAC fetal membranes is the development of an extra membrane of fetal epidermal origin that completely covers the fetus. This membrane appears attached at the mucocutaneous junctions of the lips, nose, eyes, anus, vulva or prepuce, and coronary bands. Although the precise function of this lubricated epidermal membrane is unknown, it has been suggested that it would facilitate delivery.

Parturition Almost 100% of the parturitions occur during the early hours of the day. No births occur between 17.00 and 04.00 hours, when temperatures are low throughout the year in the puna. This is an additional adaptation of these animals to the severe Andean environment. Surprisingly, SACs do not lick their offspring or eat the placenta. Alpacas and llamas are excellent mothers and rarely abandon their newborn, even when the nutritional status of the adult is poor. Symptoms of approaching parturition are minimal and not easy to detect.

Chapter 14  •  Anatomy and Physiology of Reproduction in the Female Llama and Alpaca Little attention has been paid to the endocrinologic events related to the parturition process. In most mammalian species, the fetus dominates the mechanism stimulating the onset of parturition. A significant increase in the fetal plasma concentration of cortisol occurs during the final stages of gestation in sheep, goats, cattle, and pigs. In llamas and alpacas, high cortisol concentrations are recorded only during the day of parturition.50 The role of cortisol in the induction of parturition in llamas and alpacas remains undetermined. The secretory pattern reported in this study resembles that observed in the mare, but additional studies are required to confirm and define such a role.50 In addition, massive release of PGF2α is observed during the parturition day in both species. Even more, mean PGF2α metabolite concentrations during the parturition day are significantly higher in samples collected in the morning than in the sample collected in the afternoon. This finding suggests that in llamas and alpacas, as in the mare, the second stage of labor is accompanied by an explosive rise in PGF2α con­ centrations as the newborn passes through the cervix and the vagina. Body weight of crias at birth in their natural habitat ranges from 6.9 to 8.4 kg and 9 to 14 kg in alpacas and llamas, respectively. In general, cria body weight increases with the age of the dam, peaking with 9-year-old dams, and then begins to decline. Less than 6% of perinatal mortality has been reported, and this trait is also associated with the weight of the cria at birth and the age of the parturient dam.56

Early Postpartum Period Uterine weight decreases to one fifth of that recorded at parturition by day 5 after parturition. Uterine involution is completed about 15 days after parturition in the alpaca. Even though females show receptivity as early as 24 hours after parturition, they are not fertile at this time, since uterine involution has not been completed and ovulatory follicles are not present. It is recommended that females not be mated earlier than 15 to 20 days after parturition to obtain high fertility rates.32 Fertility increases form 30% to 70% from postpartum day 10 to day 30.57 Like in mares and sows, which are species with similar placentation, prostaglandin concentrations return to basal concentrations 1 to 2 days after parturition in llamas and alpacas. Conversely, prolonged release of PGF2α during several weeks after parturition has been reported in cows, water buffalos, goats, and ewes. All these ruminants have a similar type of cotyledonary placenta. The physiologic significance of this difference in postpartum prostaglandin release between species is not fully understood, but some relationship to type of placentation might exist. Estradiol-17β plasma concentrations continue to decline after parturition until basal concentrations are registered within the following 2 days. A significant rise in estradiol-17β concentrations, indicative of follicular development is registered within 7to 10 days after parturition in llamas and alpacas. Thus, follicle growth has been observed to resume as early as 3 to 4 days after parturition, and a single ovulatory follicle develops by the time of estradiol-17β rise.50 This observation correlates well with the fact that SACs usually become pregnant within the first 2 to 3 weeks after parturition. Similarly, estrone sulphate concentrations accurately reflect ovarian follicle development in the postpartum period in llamas.46,49 If

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the long gestation period registered in camelids is taken into account, these peculiarities explain how llama and alpaca females are able to mate soon after parturition and an offspring per year is possible.

REFERENCES 1. Fowler ME, Bravo PW: Reproduction. In Fowler M, editor: Medicine and surgery of South American camelids: llama, alpaca, vicuna, guanaco, ed 2, Ames, IO, 1998, Iowa State Univeristy Press (pp 381-429). 2. Bourke DA, et al: Ultrasonography as an aid to controlled breeding in the llama (Lama glama), Vet Rec 130:424-428, 1992. 3. Adams GP, et al: In situ morphologic dynamics of ovaries, uterus and cervix in llamas, Biol Reprod 41:551-558, 1989. 4. Vaughan JL, Tibary A: Reproduction in female South American camelids: a review and clinical observations, Small Rumin Res 61:259-281, 2006. 5. Del Campo MR, et al: The application of new reproductive technologies to South American camelids, Theriogenology 43:21-30, 1995. 6. Vaughan JL: Ovarian function in South American camelids (alpacas, llamas, vicunas, guanacos), Anim Reprod Sci 124: 237-243, 2011. 7. San-Martin M, et al: Aspects of reproduction in the alpaca, J Reprod Fertil 16:395-399, 1968. 8. Bravo PW, Sumar J: Laparoscopic examination of the ovarian activity in alpacas, Anim Reprod Sci 21:271-281, 1989. 9. Chaves MG, et al: Ovarian follicular wave pattern and the effect of exogenous progesterone on follicular activity in non-mated llamas, Anim Reprod Sci 69:37-46, 2002. 10. Adams GP, et al: Effects of lactational and reproductive status on ovarian follicular waves in llamas (Lama glama), J Reprod Fertil 90:535545, 1990. 11. Vaughan JL, et al: Ovarian follicular wave characteristics in alpacas, Anim Reprod Sci 80:353-361, 2004. 12. Mirayaga MH, et al: Follicular activity and hormonal secretory profile in vicuna (Vicugna vicugna), Theriogenology 61:663-671, 2004. 13. Bravo PW, et al: Ovarian follicular dynamics in the llama, Biol Reprod 43:579-585, 1990. 14. Bravo PW, et al: Ovarian and endocrine patterns associated with reproductive abnormalities in llamas and alpacas, JAVMA 202:268272, 1993. 15. Adams GP, et al: Form and function of the corpus luteum in llamas, Anim Reprod Sci 24:127-138, 1991. 16. Tibary A, Vaughan JL: Reproductive physiology and infertility in male South American camelids: a review and clinical observations, Small Rumin Res 61:283-398, 2006. 17. England BG, et al: Oestrous and mating behaviour in the llama (Llama glama), Anim Behav 19:722-726, 1971. 18. Novoa C: Incremento reproductivo en alpacas en explotaciones comerciales mediante método de empadre alternativo, Revista de Investigaciones Pecuarias, Peru 2:191-193, 1973. 19. Fernandez-Saga S, et al: Effect of different mating stimuli on induction of ovulation in the alpaca, J Reprod Fertil 22:261-267, 1970. 20. Chen BX: Semen-induced ovulation in the Bactrian camel (Camelus bactrianus), J Reprod Fertil 73:335-339, 1985. 21. Pan G, et al: Isolation and purification of the ovulation-inducing factor from seminal plasma in the Bactrian camel (Camelus bactrianus), Theriogenology 55:1863-1879, 2001. 22. Paolicchi F, et al: Biological activity of the seminal plasma of alpacas, stimulus for the production of LH by pituitary cells, Anim Reprod Sci 54:203-210, 1999. 23. Adams GP, et al: Ovulation-inducing factor in the seminal plasma of alpacas and llamas, Biol Reprod 73:452-457, 2005. 24. Ratto MH, et al: Ovulation-inducing factor: a protein component of llama seminal plasma, Reprod Biol Endocrinol 8:44, 2010. 25. Kershaw-Young CM, et al: ß-nerve frowth factor is a mjor component of alpaca seminal plasma and induced ovulation in female alpacas, Reprod Fert and Devel 24:1093-1097, 2012. 26. Ratto MH, et al: The nerve of ovulation-inducing factor in semen, Proceedings of the National Academy of Sciences 109:15042-15047, 2012. 27. Aba MA, et al: Control of ovarian activity in Llamas (Lama glama) with medroxyprogesterone acetate, Reprod Domest Anim 34:471-476, 1999. 28. Bravo PW, et al: Pituitary response to repeated copulation and/or gonadotropin-releasing hormone administration in llamas and alpacas, Biol Reprod 47:884-888, 1992.

29. Bravo PW, et al: The effect of ovarian follicle size on pituitary and ovarian responses to copulation in domesticated South American camelids, Biol Reprod 45:553-559, 1991. 30. Aba MA, et al: Levels of progesterone and changes in prostaglandin F-2 alpha release during luteolysis and early pregnancy in llamas and the effect of treatment with flunixin meglumine, Anim Reprod Sci 59:87-97, 2000. 31. Ratto M, et al: Comparison of the effect of natural mating, LH, and GnRH on interval to ovulation and luteal function in llamas, Anim Reprod Sci 91:299-306, 2006. 32. Bravo PW: Reproductive endocrinology of llamas and alpacas, Vet Clin North Am: Food Anim Pract 10:265-279, 1994. 33. Sumar JB: Reproduction in llamas and alpacas, Anim Reprod Sci 42:405-415, 1996. 34. Fernandez Baca S, et al: Relación entre la ubicación del cuerpo lúteo y la localización del embrión en la alpaca, Revista de Investigaciones Pecuarias, Peru 2:131-135, 1973. 35. Sumar J: Removal of the ovaries or ablation of the corpus luteum and its effect on the maintenance of gestation in the alpaca and llama, Acta Veterinaria Scandinavica 83:133-141, 1988. 36. Aba M, et al: Endocrine changes after mating in pregnant and nonpregnant llamas and alpacas, Acta Vet Scand 36:489-498, 1995. 37. Bianchi CP: Estudios sobre el control endocrino de la luteólisis y reconocimiento materno de la preñez en llamas (Lama glama) (doctoral thesis), Universidad Buenos Aires, Argentina, 2011, Nacional del Centro de la Provincia de Buenos Aires. 38. Bianchi CP, et al: Uterine estrogen receptor alpha and progesterone receptor during the follicular and luteal phase in llamas, Anim Reprod Sci 99:117-126, 2007. 39. Fernandez-Baca S, et al: Differential luteolytic effects of right and left uterine horns in the alpaca, Biol Reprod 20:586-595, 1979. 40. Del Campo MR, et al: Vascular provisions for a local utero-ovarian cross-over pathway in New World camelids, Theriogenology 46:983991, 1996. 41. Ghazi R: Angioarchitectural studies of the utero-ovarian component in the camel (Camelus dromedarius), Reprod Fertil 61:43-46, 1981. 42. Skidmore JA, et al: Control of luteolysis in the one-humped camel (Camelus dromedarius), J Reprod Fertil 114:201-209, 1998. 43. Aba MA, et al: Endocrine changes during early pregnancy in the alpaca, Anim Reprod Sci 47:273-279, 1997.

44. Picha Y, et al: Chronology of early embryonic development and embryo terine migration in alpacas, Theriogenology 79:702-708, 2013 45. Skidmore JA: Reproduction in dromedary camels: an update, Anim Reprod Sci 2:161-171, 2005. 46. Bravo PW, et al: Urinary steroids in the periparturient and postpartum periods through early pregnancy in llamas, Theriogenology 36:267-278, 1991. 47. Powell SA, et al: Estradiol production by preimplantation blastocysts and increased serum progesterone following estradiol treatment in llamas, Anim Reprod Sci 102:66-75, 2007. 48. Skidmore JA, et al: Oestrogen synthesis by the peri-implantation conceptus of the one-humped camel (Camelus dromedarius), J Reprod Fertil 101:363-367, 1994. 49. Chipayo Y: Efecto del estradiol en el periodo de reconocimiento materno de la preñez sobre la supervivencia embrionaria en alpacas, Rev Inv Vet Perú 14:111-118, 2003. 50. Aba MA, et al: Plasma concentrations of 15-ketodihydro-PGF(2 alpha), progesterone, oestrone sulphate, oestradiol-17 beta and cortisol during late gestation, parturition and the early postpartum period in llamas and alpacas, Anim Reprod Sci 50:111-121, 1998. 51. Apichela S, et al: In Vivo and in vitro sperm interaction with oviductal epithelial cells of llama, Reprod Domest Anim 44:943-951, 2009. 52. Bravo PW, et al: Transport of spermatozoa and ova in female alpaca, Anim Reprod Sci 43:173-179, 1996. 53. Tibary A, et al: Current knwoldge and future challenges in camelid reproduction. Proceedings of the Seventh International Symposium on Reproduction in Domestic Ruminants, Wellignton, New Zealand, August 2006, Nottinghan University Press, Nottingham, UK pp297-313 2007. 54. Leon J, et al: Endocrine changes during pregnancy, parturition and the early post-partum period in the llama (Lama glama), J Reprod Fertil 88:503-511, 1990. 55. Olivera LVM, et al: Develpmental changes at the materno-embryonic interface in early pregnancy of the alpaca, Lamos pacos, Anat Embryol 207:317-331, 2003. 56. Bravo PW, et al: Cria alpaca body weight and perinatal survival in relation to age of the dam, Anim Reprod Sci 111:214-219, 2009. 57. Bravo PW, et al: Evaluation of early reproductive performance in the postpartum alpaca by progesterone concentrations, Anim Reprod Sci 39:71-77, 1995.