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Multiple Ovulation and Embryo Transfer (MOET)
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Collection of embryos in camelidae has been attempted since the late 1960's and early 1970's.(68, 69) However, the first successful non-surgical embryo transfer in camelidae with a birth of a live offspring was reported in the llama in 1985.(87) Many reports, particularly in the dromedary(7, 31, 32, 62, 63, 76) and llama,(13, 14, 17, 57, 60, 61) have since been published on different aspects of induction of multiple ovulation and embryo transfer. Embryo transfer has also been attempted in the alpaca with variable results.(82) Techniques of embryo transfer used in the dromedary have been applied successfully to the Bactrian camel in our laboratory.
Management of donors
Superovulation is a very important aspect of any MOET program. Superovulation allows a more efficient use of embryo transfer, especially in wild or valuable animals. Several hormonal treatments used to increase the number of ovulations in sheep and cattle have been adapted to the camelidae female with variable success. These include the use of exogenous gonadotropins such as eCG or FSH (7, 13, 14, 17, 32, 57, 62, 63, 76, 82) or immunization against inhibin (studies in progress).
Superovulation with FSH
FSH of ovine or porcine origin has been used for superovulation in the dromedary and llama. In the dromedary, a total dose of 20 to 30 units of oFSH is given over 6 days (two injections daily, according to protocols similar to those used in the bovine).(31, 32, 76) FSH treatment is started 2 days before and up to 1 day after completion of a 7-day course of progesterone treatment by intravaginal device (PRID).(31, 76) However, the authors did not specify whether the total dose was distributed in constant, increasing, or decreasing fashion. The results of this treatment were very low in terms of embryo recovery. In one study, out of 11 females treated, 8 did not yield any embryos, 1 gave one embryo, one gave 4 embryos, and another female gave 12 embryos. The interval from PRID removal to mating (sexual receptivity) was 7 and 8 days respectively for females treated with 20 and 30 Units of oFSH on the day of PRID removal and 4.3 and 3 days when treatment was given one day before PRID removal.(31) Embryo recovery rates were very poor and probably due to early breeding (before maturation of follicles) of these females. oFSH was also given in a single small dose (3.3 units) followed by an injection of 3500 IU of eCG and resulted in an average of 7 embryos recovered per treated female.(76) However, these results are likely due to the effect of eCG rather to FSH. In another study, ovine FSH was given twice a day (1 to 3 mg per injection) during 3 to 5 days following a 10- to 15-day course of progesterone treatment (100 mg per day during 10 to 15 days).(63) Porcine FSH (1 to 3 mg bid) given in decreasing doses over 3, 5, or 7 days after a 10- to 15-day progesterone treatment also resulted in superovulation of a dromedary female.(63) Similar results were obtained by our group using FSH from different origins, including camel FSH. The interval from treatment to development of mature follicles varies from 6 to 8 days. Superovulation can be obtained without previous progesterone treatment. However, the best results are obtained when the treated females have no follicular structures on the ovary. In some females only one ovary responds to superovulation treatment.
Superovulation with eCG
Equine Chorionic Gonadotropin (eCG or PMSG) is well known for its FSH activity and has been used by several authors to promote follicular development and superovulation in llamas, alpacas, and dromedary females. The dosage of eCG used varied from 1500 to 6000 IU in the dromedary,(7, 32, 62, 63, 93) 500 to 2000 IU in the llama, and 750 IU in the alpaca.(19, 34) eCG is generally injected in a single dose one day before, or on the day of, completion of a 5- to 15-day progesterone regime.
In the dromedary, eCG used in a single injection of 2000 IU, 2500 IU, or 4000 IU given one day before or one day after PRID removal resulted in superovulation in only 12 females out of 30 (40%). Only 5/12 (41.7%) ovulating females yielded one embryo or more. The interval from PRID removal to mating was 5 and 4.5 days respectively for females receiving 2500 and 4000 IU of eCG. This interval was one day shorter in females treated with eCG one day before removal of the PRID. Mating in this study was based on sexual receptivity which is not a good indicator of ovarian status and maturity of follicles.(31, 32, 76) In our laboratory, the interval from eCG treatment to mating based on the presence of a mature follicle (diameter at least 12 mm) is relatively constant (B days).
Follicular response (number of mature follicles) detected by rectal palpation following treatment with 1500 1U or 2000 IU of eCG was respectively 4.4. ± 3.2 (range 0 to 10) and 5.7 ± 5.4 (range 0 to 19).(7) The proportion of females with 4 or more follicles after treatment was 6/9 and 11/21 respectively for females receiving 1500 IU eCG and those receiving 2000 IU eCG. This experiment showed that response rate is very variable from one female to another with a possible relationship between response and dose of eCG. The proportion of females that did not show any follicular growth following injection of 1500 IU and 2000 IU of eCG was respectively 2/9 and 4/21.(7)
In the llama, superovulation was attempted by administration of eCG (1000 IU) following progesterone priming either in the form of natural corpus luteum (ovulation induced by hCG or GnRH injection), CIDR's, or subcutaneous implants.(17, 19) The follicular response to these treatments was variable (0 to 13 follicles) and the number of ovulations ranged from 0 to 7 with a mean number of embryos collected of 1.3 to 2.3 per donor (range 0 to 6). Follicles reached mature size (9 to 13 mm) 5 to 11 days following eCG treatment. Many females showed premature luteinization 7 to 9 days following eCG treatment. Increasing the dose of eCG to 2000 IU resulted in an increase in incidence of anovulatory follicles. In the alpaca, injection of 750 IU of eCG resulted in an average of 3.7 embryos per ovulating female.(16, 17, 19)
Superovulation with immunization against inhibin
Inhibin is a glycoprotein secreted by granulosa cells which regulates follicular growth through an inhibitory effect on FSH secretion. Immunization against inhibin results in very high levels of circulating FSH and consequently an increase in the number of recruited follicles which will continue to grow until maturation.
Inhibin has been isolated and characterized in several species including pigs, sheep, cattle, and horses. The biologically active molecule has a molecular weight of 32.000 Da and is composed of two sub-units α and ß united by disulfure bonds. Secretion of inhibin and estradiol by the growing follicle plays an important role in the regulation of FSH secretion and selection of the dominant follicle. Once the dominant follicle is selected, it continues to grow while the subordinate follicles regress due to low FSH levels. Neutralization of the endogenous inhibin produced by the follicles results in a maintenance of high levels of FSH during the periods of recruitment, growth, and maturation of the follicles which in turn results in a larger number of follicles reaching maturity. This neutralization of inhibin can be achieved by vaccination. Immunization is usually obtained by serial injection of synthetic or recombined fragments of the α- sub-unit. The immunogen is obtained by conjugating these fragments of inhibin with a larger protein such as bovine serum albumin (BSA) or Keyhole Limpet Hemocyanine (KLH). This product is then combined into an adjuvant in order to stimulate immunological response.
Preliminary experiments in the dromedary using synthetic peptide fragments from N-terminal sequence of Inhibin-α as an antigen are very encouraging. The fragment used was conjugated to ovalbumine and combined with complete or incomplete Freund adjuvant. Six female dromedaries received subcutaneous injections of the immunogen 4 times. The interval between injections was 2 weeks between the first and second injections and 3 weeks thereafter. Response to immunization was monitored by regular palpation and ultrasonography and breeding took place when the size of the follicles w'as adequate (12 to 16 mm). Two out of the six treated females did not show any response to immunization whereas the other 4 showed a response after 4 injections of the immunogen. The number of embryos recovered per collection in these females ranged from 4 to 10 showing that this technique can be considered for superovulation in the dromedary female. Further studies are being conducted to determine the optimal dose and rhythm of immunization, as well as the long-term effect of this treatment on ovarian function.
Problems with superovulation in camelidae
Superovulation treatments in the camelidae female are far from being perfect. Ovulation response and embryo yield remain very variable. From our experience in the dromedary' and Bactrian camels and from other work in the dromedary, llama and alpacas, the major problems that need to be addressed are:
1) The high incidence of non-responsive females: 20 to 30% receiving superovulation treatment do not develop follicles. This is probably due in some females to immunization against eCG. In FSH-treated females, we have noticed a high incidence of premature regression of the follicles which could be due to inappropriate dosage and/or method of delivery.
2) The high incidence of follicle luteinization before breeding. This is particularly prevalent in eCG-treated females and could be due to the LFf activity of this hormone.
3) The high variability in response amongst treated females.
4) The high incidence of "over-stimulated" ovaries (Figure 11.4): In some eCG or FSH super-ovulated females, the ovaries become very large and contain many generations of follicles with different sizes. This is probably due to an individual difference in response to the hormones. Most of the females with over-stimulated ovaries do not yield any embryos, probably because of disturbance of the gamete transport or fertilization processes.
5) Dromedary camels can become refractory to superovulation with FSH and eCG. This is probably caused by immunization against these hormones. We have observed a complete arrest of ovarian activity in 5 females that have been superovulated with these hormones repeatedly during several years.
More research is needed in the area of follicular dynamics and its intra-ovarian and hypthalamo-pituitary regulation in order to address these problems.
Management of breeding
Breeding management of the donor is very important in order to determine the optimal time for breeding and induction of ovulation and consequently the time for embryo collection. Although breeding will induce ovulation in some females, most of the MOET programs will use some hormonal treatment to promote ovulation in donors.
Mating and induction of ovulation
In the dromedary, some authors rely on behavioral estrus to determine the time of mating.(76) This is probably not the best method for the management of superovulated donors because signs of estrus do not correlate well with ovarian follicular status. In order to achieve a good ovulation rate, donors should be monitored by ultrasonography and palpation throughout the superovulation treatment. The decision to breed should be made on the basis of the size of the follicles. Ultrasonographic monitoring of several females in our MOET program showed that the follicular wave appears 3 to 5 days after initiation of the superovulation treatment. The follicles at this stage are generally 3 to 5 mm in diameter and disposed around the periphery of the ovary (Figure 11.5). If the treatment has been adequate, these follicles will continue to grow at a rate of I to 2 mm per day (Figure 11.5). Breeding or insemination can be started when the follicles reach 10 mm in diameter but maximum ovulatory response is obtained if the follicles are 14 to 16 mm in diameter (Figure 11.6). Breeding should never be delayed beyond the 20 mm follicular size because of the increased incidence of luteinization or failure of ovulation at this stage (Figure 11.6). The optimal time for breeding is generally reached 6 to 8 days following superovulatory treatment. The uterus is usually very contracted and shows increased edema. In over-stimulated females, several generations of follicles may be present on the ovary which sometimes makes breeding management of these animals very difficult. Many females with partially luteinized follicles, especially in the case of superovulation with eCG, will have a flaccid uterus and may be rejected by the male.
The number of breedings per donor varies according to authors. Some authors suggest 2 breedings or inseminations at 12- to 24-hour intervals.(63, 76) In our program, a single mating or insemination is sufficient; a second breeding is done only if the copulation time at the first mating was less than 3 minutes. Although ovulation can occur in response to mating, donors should receive an injection of hCG (3000 IU) or GnRH agonist (Buserelin, 20 µg) after the first mating in order to maximize ovulation response.(7, 32, 63, 76) According to our experience, hCG treatment gives a better ovulatory response than GnRH treatment in superovulated females.
In the llama and alpaca, breeding management is similar to what has been described for the dromedary. Mating is conducted when the follicles become mature (8 to 12 mm) and all mated females receive hCG (730 to 1000 IU) or GnRH (0.1 mg) or Buserelin (8 to 20 µg).(2, 13, 15, 17, 19, 60, 61, 87) Incidence of ovulation after breeding alone, breeding + hCG, and breeding + GnRH, is respectively 80%, 90%, and 81%.(2) The main advantage of treatment with GnRH or hCG is the synchronization of ovulation. Ovulation occurs on the average 2 to 3 days after breeding, 27 ± 0.3 hours after hCG. or 28.6 ± 0.4 hours after GnRH.(2)
Verification of ovulation
Determination of the occurrence of ovulation in donors is very critical for any MOET program. Ovulation in camelidae can be detected by ultrasonography or progesterone assays both in the llama (2, 15, 17, 19, 60, 61) and in the dromedary.(7, 10, 32, 58, 59, 84, 86) Ovulation detection by ultrasonography is based on the disappearance of
follicles and/or the presence of corpora lutea (Figure 11.7). Visualization of the corpora lutea is usually difficult in the first 3 days post-ovulation but becomes easy thereafter. Plasma progesterone levels start to increase 2 to 3 days after ovulation and reach high levels (> 2 ng/ml) by day 5 after ovulation. Plasma progesterone levels are highly-correlated with the number of corpora lutea but can also be elevated in the case of luteinization.(2)
Embryo collection and evaluation
Methods for the collection of embryos from camelidae are similar to those described in other species. Surgical embryo collection after exteriorization of the uterus via laparotomy is possible in the llama, the alpaca(69) and the dromedary (Tibary and Anouassi, studies in progress). However, the use of this technique is only justified when collection of embryos at the tubal stage of development are desired. The most widely used technique for the collection of embryos from camelidae remains the non-surgical technique.
Non-surgical collection of embryos
Non-surgical collection of embryos from camelidae is done in the same manner described for the bovine species. The donor is placed in palpation stocks after sedation and the rectum is cleaned of all feces. The tail should be wrapped and the perineal area and the vulva scrubbed. Some authors suggest the use of epidural anesthesia.(44, 62, 63, 76, 87) Low epidural anesthesia is advantageous in llamas and alpacas and in young dromedaries because of the smallness of the pelvis. However, it is not necessary in large females with a well-developed pelvis, especially if they are already sedated.
Collection is made using a Foley catheter (12 to 16 gauge for llamas and 18 to 22 gauge for camels).(2, 7, 13, 14, 17, 19, 61-63, 76, 87) a stylet is inserted into the catheter to keep it from bending during manipulation. In the Bactrian and dromedary camel, the catheter-stylet set is introduced into the vagina and directed toward the cervix, then manipulated per rectum in the same manner as for the inseminating gun. In some females, passage of the cervix using this technique is difficult, especially if the cervix is large or has protruding cervical rings. In such cases, the catheter which is in a long sterile, sleeve and can be guided vaginally. The cervix is then dilated manually and the catheter is inserted the first two rings of the cervical canal. Introduction of the catheter into the uterus is completed by recto-vaginal manipulation. Passage of the cervix by rectovaginal manipulation is somewhat difficult for the inexperienced practitioner especially in older, superovulated females. This difficulty is principally due to the morphology of the external cervical rings and to the increased relaxation of the uterus in the presence of a high level of progesterone (multiple corpora lutea).
In the Bactrian and dromedary camel, we always flush each horn separately because the cervical canal relaxes during flushing and the cuff can easily slip back into the vaginal cavity causing loss of flushing media. However, other authors prefer flushing the whole uterus.(62, 63, 76, 93) Whole-uterus flushing is done by placing the catheter cuff 1 to 2 cm cranial to the internal cervical os and inflating it with 30 to 50 ml of air or PBS medium (Figure 11.8). The uterus is flushed repeatedly with 30 to 70 ml of medium (DPBS + 1% FCS + pyruvate + glucose + penicillin/streptomycin) or (DPBS + 0.2% BSA + kanamycine sulfate) for a total volume used of 1000 ml. Some authors suggest that the operator's hand should be kept in the vaginal cavity for the first 500 ml of flushing to prevent expulsion of fluid from the cervix.(62)
For individual-horn flushing the catheter should be placed in the uterine horn so that the cuff is positioned at its lower third (Figure 11.8). This is sometimes difficult to judge because the uterine horns in camelidae are separated internally by a septum that is not palpable. The cuff is inflated with air or the flushing medium in such a manner that the catheter is well anchored and can not move under the pressure of the medium. The amount of medium used for inflating the cuff ranges from 25 to 40 ml according to the size of the uterine horn. The horn is flushed 4 to 5 times by injection of 30 to 120 ml of Dubelcco's phosphate-buffered saline with 1% heat-treated fetal calf serum, which is recuperated into a cylinder by gravity. The right amount to be flushed into the uterus is judged by palpation of the distended uterine horn. Recuperation of the medium must be complete. This is aided by gentle massage of the uterine horn. The operation is repeated similarly with the other uterine horn.
In the llama, insertion of the Foley catheter into the cervix can be facilitated by using a vaginal speculum. The catheter tip is advanced through the speculum and threaded into the first two cervical rings then maintained in place by exerting some pressure while the speculum is removed gently. The introduction of the catheter into the uterus is completed by recto-vaginal manipulation as described above. Most of the authors prefer to flush both uterine horns at the same time in llamas in order to avoid the need for reinsertion and manipulation of the catheter. Thus the catheter should be brought caudal to the internal cervical os after inflation of the cuff (3 to 5 ml).(13, 14, 17, 19, 87) The uterus is flushed with 30 to 50 ml of medium (DPBS + 1% FCS + peni/strep + Fungizone + Glucose (5.5 mM) and Na Pyruvate (.33 mM)) 3 to 5 times. A total of 300 to 500 ml is used per flushing.(2, 13, 14, 17, 19, 60, 87)
The flushing medium is collected in sterile siliconized cylinders and allowed to settle at 37°C for 10 to 15 minutes and up to one hour.(7, 76, 93) After this period of sedimentation most of the medium is siphoned out, using tubing and a 75 µm embryo filtering hose until only 30 ml of medium is left in the cylinder.(7) This bottom part of the flushing medium is then poured into large, sterile, (100 x 15 mm) square petri dishes with a 13 mm grid for observation under a dissecting scope. An alternative method to siphoning for the elimination of the extra flushing medium is to collect the medium directly into an embryo filter.
Factors affecting embryo yield
Embryo recovery rate
Embryo recovery rates are highly variable in camelidae and depend on many factors such as superovulation treatment, fertility, management, collection date, and experience. Embryo recovery rates in the dromedary vary from 114% to 384% and are affected by several factors.(62, 63)
Non- superovulated donors
Collection of embryos in non-superovulated animals can be done if the number of recipients is limited. Reported recovery rates in non-superovulated llamas range from 52 to 61%.(2, 57) In the dromedary, we have obtained higher rates - 85% in single ovulators and 165% in double ovulators. This suggests that fertilization can be very high if the breeding is well-managed. The difference in recovery rate between studies on the llama and our study in the dromedary is not easily explained and could be due to a difference in fertility. Recovery rate is higher in females with double ovulation and twice that obtained in a single ovulator which suggests that double ovulation does not affect fertilization rate and survival of the embryo until day 7 post-ovulation. In camelidae of South America, the incidence of double ovulation is about 10%.(43, 47, 49, 79) The incidence of double ovulation increases to 20% in animals stimulated with hCG or GnRH.(79, 87)
Embryo recovery rates in superovulated donors are highly variable (114 to 384%).(63) This high variability could be due to many factors including failure of ovulation or abnormal luteinization, poor fertilization rate, or perturbation of embryo development and transport.
Failure of ovulation can occur if the animals are bred too early (before the follicles reach their mature size) or if there is a low LH-response following breeding. To alleviate these problems, donors should not be bred before the follicles are at least 10 mm and no more than 18 mm in diameter. Also, superovulated animals should receive an injection of hCG (3000 IU) at the time, or within 24 hours, of mating.
The type of superovulation treatment can have a great effect on the embryo recovery rate. In a retrospective study on many superovulation results, FSH-treated dromedary females yielded more embryos (417% for natural mating, 188% for artificially insemination) than eCG-treated females (227% natural mating, 69% AI).(63) The low recovery rates obtained in eCG-superovulated females can be due to an increase in spontaneous luteinization associated with the high LH activity in eCG and its long-lasting effect.(2, 62, 63) Similar effect of eCG has been observed by our group.
In the llama, an embryo recovery rate of 150% was obtained following eCG (1000 IU) treatment.(17) The proportion of treated females (n = 52) that had more than 2 ovulations (superovulated) was 57.7% and only 28.8% of the treated females gave more than 2 embryos (45.5% of the flushed females and 63.5% of the females were flushed). These results would be considered poor compared to results of MOET programs in other species. Recovery rate in the guanaco and llama was found to be inversely proportional to the number of corpora lutea. This could be due to a failure of fertilization or to an incapacity of the fimbria to collect all oocytes.(19) Number of matings seems to be an important factor in embryo recovery in superovulated llamas. In one study, the embryo recovery rate was 22% and 72% in females mated once or twice respectively.(19)
From the limited number of studies published and our own embryo transfer results we can conclude that the major factors affecting the efficiency (number of embryos) of superovulation with exogenous hormones are the time of initiation of the treatment, donor follow-up, breeding decision, and the number of ovulations obtained. Our preliminary results show that although superovulation treatment can be started any time during the follicular wave cycle, the best results are achieved when treatment is initiated in the absence of any follicles, 2 days after induction of ovulation, and after a 10- to 15- day progesterone treatment. All donors should be examined every other day to evaluate follicular growth and a breeding decision should be made based on the size of the follicles and the tone of the uterus. Ovulation response and fertilization are increased if the donors are bred when the follicles reach at least 12 mm in diameter and the tone of uterus is maximal. However, breeding of donors with low uterine tone because of the presence of luteinized follicles or even a corpus luteum has yielded embryos in our laboratory.
Effect of timing of flushing on embryo recovery rate
It is well established now that in camelidae, the embryo does not reached the uterus until day 6 or 6.5 after ovulation. Therefore any attempt to collect embryos from the uterine cavity before day 6 post-ovulation will result in a low recovery rate.(2, 7, 17, 63, 87) In practice, uterine flushing in the llama and guanaco is done 6 to 9 days after GnRH treatment.(2, 14, 17, 19, 60, 87) No embryos are recovered from the uterus of females flushed at day 5.5 or 6 post-ovulation.(2) In the dromedary, the uterus is generally flushed between 6 and 7 days after mating and treatment with hCG or GnRH.(7, 31, 62, 63, 76) However, the best embryo recovery rates are obtained when the donors are flushed on day 7 or later. In two separate studies, the recovery rates were 114%, 179%, and 267% after flushing on day 6, day 6.5, and day 7 after mating, and 60%, 157%, 175%, 200%, and 300% for collections at day 6, day 6.5, day 7, day 7.5, and day 9, respectively.(62, 63) The effect of collection date on embryo recovery rates is even more pronounced in superovulated females because of possible delay in oviductal transport and fertilization. This observation is substantiated by the high frequency of pregnancies observed in donors after embryo collection. In fact, many donors become pregnant even after two flushings at 12- to 24-hour intervals.(62, 63)
Inexperience of the operator can account for many unsuccessful flushings. In our laboratory, recovery rates on single ovulators increased from 60% to 85% with increased experience of the operator. Most inexperienced practitioners have difficulties appreciating the position of the cuff medium and can go too far into the uterine horn. A badly positioned cuff tends to slip back and fluid is lost in the vaginal cavity. Insertion of the catheter into the left horn may be difficult for left-handed palpators and vice versa. Harsh manipulation can also lead to bleeding which makes embryo identification very difficult.
Failure of ovulation results from inadequate, or lack of, LH release. In the dromedary, we have observed ovulation failure in 15 to 20% of the donors even if the criteria for breeding decision are respected.(10, 84) Ovulation failure is more common in superovulated animals and is demonstrated by the development of large anovulatory follicles.(93) Failure of the follicles to ovulate occurs even when the females are injected with hCG or GnRH suggesting that some of the follicles may lack appropriate receptors for these hormones. In some cases, follicles regress after breeding instead of ovulating. This could be due to an improper ratio between FH and FSH during the superovulatory treatment.
Poor ovulation response can also occur when breeding decision is based only on receptivity of the female. This is due to the fact that superovulated females can be receptive even in the presence of small immature follicles which will fail to ovulate. This may be the reason why some early work on embryo transfer in the dromedary achieved very low embryo recovery results.(7, 31, 32, 77) In eCG-treated females, ovulation failure can be due to early luteinization of the follicles.
Lack of fertilization
Fertilization failure is a common problem in mismanaged animals. Reduced or complete lack of fertilization has been documented in some males with low-quality semen. In one study, all females mated to a male that had low sperm concentration (< 20 million /ml) failed to produce an embryo.(62, 63) Other factors besides semen quality can be involved in poor fertilization and embryo recovery rates. We observed a case of complete lack of fertilization despite the male's normal semen concentration and motility.
Several pathological problems such as salpingitis and endometritis affect fertilization and embryo survival and reduce embryo yields in donors. In the case of endometritis, the flushing fluid is usually opaque or cloudy and contains much cellular debris and pus. Fertilization and embryo survival may also be affected by the age of the donor. In one study on the dromedary, it has been found that embryo yield is higher in donors aged 12 years or more than in younger females (285% vs. 149%). Multiparous females yielded more embryos than nulliparous females (353% vs. 121%).(62, 63) The effect of age could also be due to increased incidence of overstimulation of the ovaries in younger females.
Fertilization of ova and embryo transport can also be hindered by the presence of large anovulatory follicles. Embryo recovery rate was found to be higher in females without (220%), than in females with, anovulatory follicles (148%).(62, 63) The incidence of anovulatory follicles is 8 to 10 times higher in females superovulated randomly (43%) than in females superovulated following a 10- to 15-day progesterone treatment course (4%).(62, 63) This suggests that progesterone treatment is able to regulate the follicular wave and increases the ovulatory response of follicles.
Lower embryo-recovery rates, probably due to reduced fertilization, are also observed when donors are artificially inseminated. Recovery rates in donors bred naturally are consistently higher (299%) than in females inseminated with fresh raw (199%) or extended semen (124%).(62, 63) No study has been done on embryo-recovery rates following artificial insemination with frozen semen. The negative effect of artificial insemination on the recovery rate of embryos could be due to an inadequate stimulation of ovulation or a poor fertilization rate.
All equipment used for flushing should be verified before use. We have had complete lots of flushing catheters with a defective cuff. Most of the embryos collected from the uterus in camelidae are at the hatched blastocyst stage. This usually makes them more susceptible to sticking to rough surfaces and to cellular debris and easy to lose during manipulation.
Evaluation of embryos
Embryos recovered from the uterus in camelidae are generally at the hatched blastocyst stage.(2, 7, 17, 31, 32, 62, 63, 76, 87) The size of the embryo is highly variable at different stages post-ovulation.
In the llama, the average diameter of the embryo is 0.3 mm (range = 0.1 to 0.7 mm), 0.9 mm (range = 0.5 to 1.0 ), and 7.5 mm (range = 4.0 to 16.0) respectively at 6.0 to 6.5 days, 6.5 to 7.0 days, and 8.5 to 9.0 days post-ovulation.(2, 17) The embryo which is spherical during the first 7 to 8 days becomes elongated at 8.5 days and its length increases steadily to reach 83 mm by day 14.(2, 17)
In the dromedary, the embryo enters the uterus between 6 and 6.5 days after ovulation at the early blastocyst stage with its zona pellucida. However, these stages seem to be very short in duration and the majority of the embryos collected at day 6.5 or 7 have already hatched by day seven. In one study, 75% of the embryos collected on day 7 after mating were already at the blastocyst stage.(76) Like for the llama, the embryos recovered from the dromedary camel 7 days after mating are very variable in size and have a diameter ranging from 175 to 500 µm.(76) This variability of the stage of development is probably due to the wide spread of ovulations in superovulated animals. Hatched embryos continue to grow rapidly and become easily visible to the naked eye as they expand. They start losing their spherical form by day 8.5 or 9 post-ovulation and become folded (Figure 11.9).(7, 63)
The evaluation system used by most authors classifies the embryos into 5 grades according to their morphological characteristics and stage of development (Table 4) (Figure 11.9, 11.10, 11.11). The clinician should look for abnormalities such as extruded blastomers, signs of degeneration (dark areas), and obvious morphological anomalies such as folding or wrinkling.
|Grade of the embryo||Characteristics|
|Grade I||Excellent quality embryo. Size corresponds to the stage of collection in relation to ovulation. Before day 8 should be perfectly spherical with smooth surface|
|Grade II||Good embryo, same as above with some irrigularities of the contour and very few protruded cells|
|Grade III||Medium quality, small embryo with dark patches, irregular contour and some protruded cells|
|Grade IV||Collapsed embryos showing dark area of degeneration and many extruded cells|
|Grade V||Non transferable. Collapsed very dark embryos or embryos that are retarded, dark morula, and all stages collected from the uterus that are younger than morula and unfertilized ova|
Management of recipients
The quality of the recipient is the most important factor in the success of any embryo-transfer program. The two main aspects of selection of recipients for embryo transfer are the screening for reproductive and health problems and the preparation or synchronization with the donor.
Criteria for selection of recipients
The overall success of an embryo transfer program is judged by the delivery rate and the survival and growth rate of the offspring. Therefore, selection criteria for a recipient in an embryo transfer program should guarantee a good fertility, normal pregnancy, and normal lactation. In our embryo transfer program the success rate doubled when a strict recipient-selection program was implemented. This screening program can be summarized as follows:
1) History: The potential recipient should be young (less than 12 years of age), have had at least one normal pregnancy with a normal delivery, and be either currently pregnant or recently weaned.
2) General examination of the recipient should focus on conformation and a good body condition and on symptoms of debilitating or contagious diseases. All potential recipients should be tested for brucellosis and trypanosomiasis.
3) A complete breeding-soundness examination should be done on the potential recipient including palpation and ultrasonography of the reproductive tract, uterine culture, vaginal examination, and examination of the udder.
4) In addition to regular examination of the reproductive tract, a normal uterine biopsy should be obtained from any female with incomplete history.
Synchronization with donors
It is now well established that the chances of survival of an embryo depend largely on the endocrinological environment in the recipient. This is why synchronization of the reproductive cycle between the donor and the recipient is very critical. Embryo-transfer results in the llama and dromedary suggest that the best recipient should have ovulated on the same day as the donor or up to 48 hours after.(7, 14, 17, 19. 63, 71)
Synchronization of the uterine environment between the donor and the recipient can be obtained by synchronization of ovulation or by complete preparation with hormonal therapy.
Synchronization of follicular development and ovulation
Synchronization of ovulation between the donor and recipient can be approached using two methods: selection of recipients from a random group or preparation of recipients in such a manner that follicular development is synchronized with that of the donor.
In the first technique, a group of cycling recipients is examined 24 hours after the donor is bred and all females that have a mature follicle and uterine tone are treated with GnRH or hCG. This method of selection of recipients is, however, time consuming and can only be used if the number of donors is limited or if the donor is not superovulated.(7, 14, 17, 19)
Synchronization of follicular development in donors and recipients has been attempted using progestagen with variable degrees of success.(2, 19, 32, 63, 76) Synchronization of recipients with Progesterone Releasing Intravaginal Devices (PRID's) (7 days) has been attempted in the dromedary but was not successful.(32) Only 39% and 28% of the females responded by ovulation after receiving respectively hCG (3000 IU) and Buserelin (20µg) following the PRID's removal. A large proportion of the females had high levels of progesterone before treatment with hCG or GnRH analogue and were considered as having ovulated spontaneously as a response to progesterone priming. This high progesterone in non-treated animals is not due to spontaneous ovulation as suggested by the authors but rather to the presence of luteinized anovulatory follicles which develop in 30 to 40% of non-bred dromedary females.
The best results of synchronization of ovulation are obtained when recipients are induced to ovulate with hCG (3000 or 5000 IU) or Buserelin (20 µg) following a treatment combining progesterone and eCG. The recipients are treated daily with progesterone (100 mg/ day) for 10 to 15 days followed by administration of 1500 to 2500 IU of eCG. Progesterone treatment is scheduled to end on the day of injection of gonadotropin in the donor.(63) The eCG treatment guarantees the presence of mature follicles in the recipient at the same time or 24 to 48 hours after the donor. However, the dose of eCG and ovulatory response of the recipient significantly affects the pregnancy results. Pregnancy rate is lower in recipients with 1, 2, or more than 6 corpora lutea compared to recipients with 3 to 6 corpora lutea.(63) Therefore, a high response of recipients to eCG is not desirable because it may cause early regression of the corpora lutea.
Preparation of recipients with progesterone
Synchronization between the embryo and the uterus can be obtained by progesterone therapy without induction of ovulation.(76) Progesterone (100 mg) is given daily starting 2 days after mating of the donor. However, because there is no corpus luteum, progesterone treatment should be continued throughout pregnancy.
Bilaterally ovariectomized females can also be used as recipients. The females are treated for two days with estradiol 17ß (40 mg/day) followed by daily injections of progesterone (100 mg). This treatment has resulted in a 30% pregnancy rate in our laboratory but has been abandoned because of the need for daily progesterone treatment throughout pregnancy.(86)
Screening of recipients
All recipients should be screened on the day of transfer to ascertain that ovulation has occurred and that a mature corpus luteum is present. This screening can be done either by determination of plasma progesterone level or by ultrasonographic visualization of the corpus luteum.
Plasma progesterone level equal or superior to 2 ng/ml is a good indicator of the presence of a mature corpus luteum. This technique has been used successfully in our laboratory for the screening of recipients 7 days after induction of ovulation. However, this technique can yield poor results if the recipients are screened on the 5th and 6th day post-ovulation because progesterone levels are not high enough in some females at this stage even if ovulation has occurred.
Ultrasonographic detection of the corpus luteum remains the most accurate and rapid technique for the detection of the corpus luteum if done by an experienced individual (cf. Ultrasonography). In addition, this technique allows the screening of recipients for pathological problems such as the presence of fluid in the uterine cavity.
In the estrogen-progesterone-prepared recipient, selection is based on the echotexture and tone of the uterus. Ideal recipients should have a homogenous uterine echotexture with absence of fluid or edema and a relaxed uterus.
Transfer of embryos
Transfer of embryos into the recipient can be done using a surgical or a non-surgical technique.
Surgical embryo transfer
Surgical embryo transfer in the dromedary and Bactrian camels is done via a left flank laparotomy. The embryo is transferred into the uterine cavity through a puncture made in the exteriorized horn by a Pasteur pipette.(63) In the dromedary, it is helpful to make a small incision of the serosa first before perforating the uterine muscular layer and depositing the embryo. However, this technique cannot be used in young or primiparous animals because the uterine horn is too short and difficult to exteriorize.
In llamas and alpacas, surgical transfer is done via flank or midline laparotomy (cf. Surgery in the female). The uterine horn is exposed through the surgical opening. Although some authors have suggested that transfer should be done into the uterine horn ipsilateral to the ovary bearing the corpus luteum, this is not necessary. In fact, there might be an advantage in transferring embryos into the left horn because of its role in systemic release of prostaglandin F2α (cf. Female physiology). In addition, the left uterine horn is usually easier to exteriorize because it is larger than the right horn. The midline approach is more practical for the exteriorization of the uterine horn in small primiparous females.
Other methods used in sheep and goats, such as endoscopy, could offer a better approach for transfer into small llamas and alpacas. These techniques will be very suitable for the vicuña and guanaco because they have less complications.
The non-surgical technique for embryo transfer consists of placing the embryo directly into the uterine lumen through the cervix using a regular bovine insemination gun. The embryo is loaded into a 0.25 ml or 0.5 ml sterile plastic straw and placed in the gun for transfer. The inseminating gun is first covered by a sterile sheath with a side opening then a second plastic sanitary sheath.(7, 13, 14, 17, 62, 63, 76, 87)
The recipient is prepared in the same manner described for embryo collection. Some authors suggest the use of epidural anesthesia. The inseminating gun is introduced into the vagina and guided towards the cervix. The sanitary sheath is perforated after passage of the first cervical ring and the gun is further inserted into one of the uterine horns. Cervical passage and uterine deposition of the embryo should be done as quickly as possible to avoid excessive irritation of the cervix and uterine mucosa which may cause prostaglandin F2 alpha release and corpus luteum demise.
Alternatively, insertion of the transfer pipette can be guided vaginally with a sterile gloved hand.(62 63, 76) This technique is preferred because cervical manipulation per rectum is difficult.
Some authors have suggested treatment of the recipient with non-steroidal anti-inflammatory drugs (flunixin meglumine 500 mg, IV) to prevent liberation of prostaglandin from the uterus as a result of cervical manipulation.(76) In addition, recipients receive antibiotics for the prevention of uterine infection. For progesterone-prepared recipients, administration of progesterone (100 mg/day IM) should be continued until the end of pregnancy. Attempts to replace progesterone treatment by synthetic progesterone silastic implants results in abortion.(76) Pregnancy diagnosis in recipients can be done as early as 8 days after transfer. However, in order to increase the accuracy of diagnosis, it should be delayed until two weeks after transfer. Verification of the presence of a normal corpus luteum of pregnancy is very important for the decision to stop or continue progesterone therapy. If there is no corpus luteum, maintenance of pregnancy will require daily injection of progesterone (100 mg) until parturition.(76) Recipients maintained pregnant with exogenous progesterone should be checked periodically for abnormalities of pregnancy such as pyometra and maceration. In our laboratory, we have found that a non-negligible proportion of recipients maintained pregnant with exogenous progesterone experience dystocia due to insufficient relaxation of the cervix.
Pregnancy results obtained with embryo transfer in camelidae
Depending on the study, pregnancy rates obtained following embryo transfer in the dromedary vary from 8 to 50%.(63, 76) Pregnancy rates are significantly affected by season, quality of recipients, and synchronization between donors and recipients. However, factors such as side of transfer, method of transfer, and age of the embryo do not have any effect on pregnancy rates.(62, 63) Our results in non-surgical embryo transfer are presented in Tables 5 & 6.
In the llama, alpaca, and guanaco, pregnancies and live births have been obtained after transfer of 6-to 7-day-old embryos (0.4 to 5 mm ).(13, 14, 17-19, 60, 69, 87) However, evaluation of the effect of different factors on pregnancy rates cannot be conducted at this time because of the small number of transfers realized in this species. Pregnancies were obtained in recipients that had ovulated either on the same day as, or one day before or after, the donor. Transfer of 2 embryos into the same recipient resulted in a single pregnancy.(2, 13, 14, 17, 19, 60, 87)
|Average embryo per female||1.16|
|Pregnancy rate after transfer||42%|
|Interval between collection and breeding (days)||4.1|
|Interval between two collections (days)||21|
|Number of collections per season||8|
|% responding to superovulation||70||70||80|
|Average number of ovulations (corpora lutea estimated by ultrasonography||2 to 25||2 to 25||2 to 8|
|Number of collected embryos||0 to 21||0 to 19||1 to 8|
|Stage of embryos at collection:|
|% Hatched blastocysts||90||90||100|
|% Morula or early blastocyst||8||6||-|
|Pregnancy rate after transfer||42||38||49|
The conjugated fragment of inhibin used in this study was provided by M. Blanc (INRA, France)
Effect of the quality of embryo
Pregnancy and parturition rates following embryo transfer can be affected by several factors. Data available on embryo transfer in camelidae is still very limited and therefore the effect of different factors on success rate have not been fully investigated. Most what is known today on this aspect of embryo transfer concerns the dromedary camel.(62, 63)
The relationship between embryo quality and pregnancy rate is difficult to establish because of the difficulty in evaluating the hatched embryo. In a study where embryos were graded from 1 to 5, pregnancy rates were higher for embryos Grade 1 than those grade 2 or more. Retrospective analysis on the morphological characteristics of each embryo transferred and the pregnancy outcome suggests that embryos can only be classified into 3 major categories: good-quality transferable embryos, medium-quality transferable embryos, and non-transferable embryos. The first category includes all hatched embryos with normal development according to the date of collection. These embryos can be round or collapsed but should have a good brilliance and not show many-extruded cells. The medium-quality embryos present several types of anomalies, including loss of brilliance (become darker), poor outline definition, or tears. Non-transferable embryos include all embryos that are arrested, have substantial loss of cellular material, or are in a very advanced stage of degeneration as revealed by their general morphology or color. It is interesting to note that in our experience none of the non-hatched embryos resulted in a pregnancy even though they resembled a normal morula. It is possible that these embryos were arrested or that the dromedary embryo is very sensitive at this stage. The manipulation of morulae may hinder the hatching process. Two embryos recovered while undergoing the process of hatching resulted in pregnancies after transfer. Similarly, two other studies on the dromedary showed that pregnancy rates achieved after transfer of early blastocysts or morulae were lower (26%) than those obtained after transfer of hatched embryos (35%).(62, 63) Although these authors obtained no pregnancies after transfer of collapsed embryos, our results suggest that the shape of the embryo does not reflect the health status of the embryo as much as its brilliance does. In our studies, the quality and survival of the embryo after transfer seem to be negatively correlated with the number of embryos in the same flushing. However, other authors have reported higher pregnancy rates following transfer of embryos from collections yielding more than 5 embryos than from collections yielding less than 5 embryos (36% vs. 26%).(62, 63) In our experience, pregnancy rates obtained from non-superovulated females are invariably of excellent quality and yield better pregnancy results than embryos from superovulated females. The decision to transfer an embryo or not based on its apparent quality should be weighed against the value of the embryo (embryo from very valuable females). Survival of the embryo and maintenance of pregnancy is possible in a wide variety of morphological appearances (Figure 11.11).
Effect of quality of recipient
The fertility of the recipient is probably the most important factor in the success of embryo transfer in any species. We have noticed a tremendous progress in pregnancy and birth rates after adopting a strict recipient-selection program based on uterine culture and biopsy. Similarly, other authors have reported that recipient fertility, age, and parity have a significant effect on pregnancy rates following embryo transfer in the dromedary. (62, 63) Recipients that are 11 years old or younger have a pregnancy rate 10 percentage points higher than recipients that are more than 11 years of age. In addition, the pregnancy rate in maiden or primiparous recipients was almost double that of multiparous recipients.(62, 63) The effect of age and parity is certainly due to the increased reproductive problems with advancing age and number of parturitions. Pregnancy rate is also affected by the number of coipora lutea in the recipient. Recipients with 3 to 6 corpora lutea have a better chance of maintaining pregnancy than those with 1, 2, or more than 6 corpora lutea.(62, 63)
Effect of synchronization
Pregnancy rates are higher if the embryos are transferred into recipients that have ovulated 1 to 2 days after the donor than those that have ovulated on the same day or earlier. In one study, the highest pregnancy rates were achieved when an embryo was transferred into the uterus of a recipient that had ovulated 1.5 days after the donor.(63) Transfer of embryos into recipients that had ovulated one day before the donor resulted in no pregnancies or a very low (10%) pregnancy rate.(63) Serial examination of the recipients every 3 days after transfer allowed us to show that the main reason for this low pregnancy rate in the recipients that had ovulated before the donor was the early demise of the corpus luteum. The luteolysis is due to the nature of the luteal phase and probably to prostaglandin F2 alpha release caused by the manipulation of the cervix and/or irritation of the uterine mucosa. In addition, pregnancy rates in recipients that ovulated 1 to 2 days before the donor and continued to receive daily injections of progesterone were similar to those achieved in recipients that ovulated 1 to 2 days after the donor. However, the progesterone treatment has to be continued throughout pregnancy which presents many disadvantages. In addition to the fact that daily administration of progesterone is time-consuming, this treatment presents some health risks such as development of pyometra, complications at birth, and inadequate udder preparation (Anouassi and Tibary, unpublished). Although some authors have reported that dromedary females can give birth spontaneously even when receiving progesterone,(76) we have observed various problems in our facilities including incomplete dilation of the cervix, premature placental separation, and inadequate milk production in recipients receiving progesterone.
To avoid the loss of pregnancy due to luteolysis we maintain the recipient under exogenous progesterone treatment until the day of pregnancy diagnosis. At pregnancy diagnosis, the presence and size of the corpus luteum are evaluated. If at 15 days post-transfer the corpus luteum is 20 mm or more in diameter, progesterone therapy is stopped. If the corpus luteum is absent or shows signs of regression, the pregnancy is maintained with exogenous progesterone and the recipient receives a dose of eCG (1500 to 2000 IU) in order to promote development of a new follicular wave. All recipients receiving eCG are re-examined a week later and given hCG (3000 IU) if follicles of more than 12 mm in diameter are present. This technique allows us to "create" new corpora lutea of pregnancy in many recipients that otherwise would have lost their pregnancy because of the demise of the initial corpus luteum. This approach prevents the need of maintaining pregnancy in recipients with no corpus luteum by daily progesterone treatment which is very labor-intensive and very expensive (Tibary and Anouassi, work in progress).
Effect of method of transfer
It is usually expected to have higher pregnancy rates following surgical transfer than following cervical transfer. This does not necessary hold true for the dromedary where comparable (30 to 40%) pregnancy rates are achieved with both.(7, 62, 63, 76)
Effect of side of transfer
Many authors have suggested that transfer of embryos into the left hom would be more advantageous than transfer into the right hom.(76, 87) This is based on the fact that nearly all pregnancies are carried in the left hom in camelidae. In addition, there is a differential release of PGF2 alpha between the left and right uterine hom. Luteolytic activity from the right hom is localized but that of the left horn is both systemic and local.(43, 48) Therefore, placing the embryo in the left horn will inhibit prostaglandin F2 alpha release into the blood stream and prevent luteolysis even if the corpus luteum is located on the right ovary. To avoid the luteolytic effect of the left horn, some authors prefer using only recipients that ovulate on the left side.(87) For the dromedary, our work and that of others does not show a significant effect of the side of transfer in relation to the side of ovulation. Pregnancy rate and incidence of luteolysis following transfer is similar for recipients transferred into the left hom or the right hom whether the corpus luteum is on the right or the left side.(62, 63)
Effect of embryo treatment
There are no available studies on the best conditions (medium, temperature) for the storage of camelidae embryos until transfer. Most authors working in this field use the guidelines proposed for the bovine and equine embryos. The embryos are stored for less than 3 hours at room temperature in small petri dishes containing an enriched preservation medium which is different from the collection medium. In addition, we cover the petri dishes to prevent direct exposure of the embryos to light. There is definitely a need for research concerning the optimal manipulation condition for the fresh camelidae embryos.
Embryo freezing was tested in our laboratory and by others but pregnancy rates following transfer of these embryos are still somewhat low (10 to 15%).(62) The freezing technique used is similar to that described for the bovine and equine species as described later in this chapter.
Effect of the technician
Studies have failed to show any significant effect of the experience of the technician on pregnancy rates following cervical transfer.(63) In our laboratory, the pregnancy rate varies from 22% to 68% according to the expertise of the technician. This is probably due the fact that we use the recto-vaginal method to bypass the cervix which is more difficult to learn than the direct-vaginal method. However, the direct-vaginal method is not always possible especially in young or nulliparous females or if the hand of the technician is too large to penetrate the vagina.
Effect of season
Although the female camelidae is capable of ovarian activity throughout the year, especially if well fed, pregnancy rates are nil if embryo transfer is conducted during the hottest months of the year.(62, 63) We have observed similar trends in our center with a reduction of pregnancy rates and an increase in early embryonic death both in females that are transferred and those that are inseminated from May to September. Such breeding dates should be avoided not only because of the poor fertility but also because of the poor growth of young born during these months.
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