Assisted Reproduction and Embryo Manipulation

Assisted fertilization is a relatively new field in veterinary medicine that is gaining a lot of interest as a research tool and as an alternative reproduction method for endangered breeds or species, as well as for the treatment of some cases of infertility in valuable animals. Assisted-reproduction techniques include in vitro fertilization (IVF), intra-fallopian gamete transfer (GIFT), and some manipulations of the embryo or gametes permitting increase of number of offspring in vitro (splitting, cloning). We present a discussion of the advantages and feasibility of these techniques in camelidae although very limited research has been published in these species.

In vitro fertilization

In vitro fertilization is still at the experimental stage in camelidae. Embryos have been produced in vitro from llamas. However, there are still no reported live offspring from embryos produced in vitro. It is obvious that there is lack of scientific data on in vitro fertilization since most of the techniques used for the preparation of semen and oocytes are adapted from work done in the bovine species. Success rates of in vitro fertilization are still very low and no embryo has been obtained so far with ejaculated semen. However, because of its multiple advantages, this field is expected to develop very quickly and reach commercial feasibility within the next five years.

In vitro fertilization offers several scientific, production, and medical advantages which can be summarized as follows:
*    Enables "treatment" of infertility in valuable animals. In the female, in vitro fertilization can be used to produce embryos from animals suffering from occluded oviducts and severe uterine or cervical pathologies. Also, males with very low semen quality can be reproduced using techniques such as sperm microinjection.
*    Allows scientists to have access to embryo development stages that would otherwise be very difficult to obtain in the live animal. Such techniques require young, oviductal-stage embryos so that freezing, splitting and cloning would be possible.
*    Answers the critical need for reliable methods of evaluation of fertilizing ability of semen
*    Offers a model for the study of fertilization and embryonic development
*    Allows the use of genetic material from females that have died if a program of oocyte preservation is implemented. Oocytes can be collected from animals that recently died or have terminal diseases and be stored in a frozen state until use. This is particularly interesting for the preservation of genetic material from zoo or wild animals.
*    Permits use of sexed semen for the production of embryos with a predetermined sex

The success of in vitro fertilization depends on several factors such as the origin and preparation of oocytes and semen, conditions of fertilization, and embryo development. In order for the oocytes and semen to be able to complete fertilization they have to undergo biochemical and morphological changes called maturation for the oocytes, and capacitation and acrosome reaction for the spermatozoa.

Oocyte collection and maturation

There are two sets of methods available for the collection of oocytes that may be applied to the camelidae: collection of oocytes in vivo and collection of oocytes after ablation of the ovary. The in vivo methods are the most desirable if a female is to be used continuously in an IVF program. However, collection of oocytes after ovariectomy or postmortem is probably the most efficient technique for experimental work on IVF.

In vivo methods for the collection of oocytes

Collection of oocytes from the live animal can be done surgically or via an ultrasound-guided aspiration of the follicles. Surgical collection of complete follicles has been practiced by our group in eCG- or FSH-superovulated females. The ovary is approached via a laparotomy and all mature follicles (10 to 15 mm) are removed from the surface of the ovary and dissected in the laboratory. This technique works relatively well if the animals are not overstimulated but is not practical for repeated recovery of oocytes from the same female. Oocytes are removed from the follicles under a dissecting microscope. The recovered oocytes are usually in an advanced stage of maturation, especially if the technique is performed 12 to 24 hours after induction of ovulation with hCG. However, recovery rate can be very low after this treatment because the follicles become very fragile and bleed easily.

The best in vivo oocyte collection technique is transvaginal ultrasound-guided follicular aspiration. The female is sedated and is prepared, as for embryo collection, with a low epidural anesthesia. A vaginal probe (curvilinear or sector probe, 5 or 7.5 MhZ) attached to a long handle (30 to 45 cm) is inserted into the vagina to the fomic. The ovaries are manipulated rectally and oriented towards the probe. Once the follicles are identified and fixed, a long aspiration needle is introduced through the probe, advanced slowly through the vaginal wall, and the follicular content is aspirated using an aspiration pump. This technique requires minimal manipulation and provides a recovery rate of 40 to 50%. In the llama and alpaca, follicular aspiration can be accomplished by laparoscopy.

Collection of oocytes after ovariectomy or post-mortem

Most of the research on IVF uses Cumulus-Oocyte-Complexes collected directly from ovaries taken after slaughter or after ovariectomy. The ovaries are washed in saline and placed in a thermos at 38°C and transported within 30 minutes to the laboratory where they are rinsed and placed in saline plus antibiotics. Cumulus-Oocyte-Complexes (COC) are collected after mincing the ovary with a razor blade and observation under a dissecting microscope. The recovered COC are washed in a modified Tyrode's solution TALP-HEPES containing: 114 mM NaCl, 3.1 mM KCl, 2.0 mM NaH₂C0₃, 0.3 mM NaH₂PO₄, 10 mM sodium lactate, 2 mM CaCl₂, 0.5 mM MgCl₂, and 10 mM HEPES supplemented with 3 mg/ml of BSA fraction V, 25 p.M sodium pyruvate and 50 µg/ml gentamycine sulphate. This technique yields on the average 6.4 oocytes per llama.^(34-36)

Oocyte evaluation

The COC are examined to determine their stage of maturation, especially the expansion of the cumulus cells. Oocytes collected just before ovulation usually have a very expanded and sticky cumulus cell layer. The COC obtained by mincing of the ovaries come from a heterogeneous population of follicles and are generally at different stages of maturation. Selection of the oocyte complexes before incubation is very important because it has been found in the llama that 17 to 52% of the oocytes are degenerated after recovery from follicles.^(34-36)

In vitro maturation

In most mammalians, the oocyte remains in the diploten stage of the meiotic prophase during all of the follicular development. As ovulation nears, the oocyte emerges from this stage of dormancy to continue its meiosis. This reactivation of the meiotic division is called oocyte maturation.

Two criteria are often used to describe the stages of oocyte maturation: cytoplasmic and nuclear maturation. Cytoplasmic maturation is represented by the migration of the cortical granules towards the oolema (vitelline membrane) and acquisition of microvilli at the surface of the cells. Nuclear maturation is represented by the appearance of a germinal vesicle and evidence of meiotic activity (metaphase and anaphase I and metaphase II).

In vitro maturation of COC from the llama was accomplished by incubation at 38.5°C under 5% C0₂ in air with high humidity in tissue culture medium TCM 199 supplemented with 0.5 µg/ml oFSH, 5 µg/ml oLH, 1 µg/ml estradiol 17-α, 25 µM pyruvate, and 10% donor bovine steer serum.^{(35, 36)} Under these conditions of incubation, 62% of the oocytes are in the Metaphase II after 36 hours of culture.

Sperm preparation

At the end of spermatogenesis, spermatozoa are not yet capable of fertilization. This characteristic is acquired in the epididymis. This process, called "epididymal maturation", takes place during epididymal transit and is completed when the spermatozoa reaches the tail of the epididymis.

One of the major characteristics of mature spermatozoa is the acquisition of motility which results from biochemical changes in the plasma membrane. These modifications concern mainly alterations of the glycoproteins and lipids of the membrane, especially the integration cholesterol. Even after this maturation, spermatozoa do not become capable of fertilization until they have been exposed to the female genital tract for some time. Once completed, this process (called capacitation) allows spermatozoa to become capable of attachment to the zona pellucida of an oocyte. The major result of the capacitation process is the alteration or elimination of substances that were adsorbed to, or integrated into, the plasma membrane of spermatozoa during their epididymal transit or after contact with seminal plasma. These modifications prepare the sperm cell to undergo other changes, especially at the level of the acrosome (acrosome reaction) which are the final changes needed for the sperm cell to penetrate the oocyte envelopes.

In most species, capacitation of spermatozoa takes place in the distal part of the isthmus of the uterine tube. Mechanisms controlling capacitation at this level are not fully understood. Several substances present in the oviduct have been identified as capacitating factors and include glucosaminoglycans, taurine, hypotaurine-α amylase,    glucuronidase, proteinase, neuraminidase, arylsulfatase, fucosidase, and acetylhexosaminidase.

It has been known for some 30 years now that the changes brought about by capacitation can be obtained in vitro in chemically defined media. In camelidae, there are no studies on capacitation and acrosome reaction in ejaculated semen. The only studies on IVF in the llama used semen from the cauda epididymis which are easier to capacitate than ejaculated spermatozoa. Epididymal sperm is washed by centrifugation in TALP medium composed of 100 mM NaCL, 3.1 mM KCl, 25 mM NaHCO₃, 0.3 mM Na₂PO₄ 21.6 mM sodium lactate, 2.0 mM CaCl₂, 0.4 mM MgCl₂, 10.0 mM HEPES, 1.0 mM sodium pyruvate, 3 mg/ml BSA, and 50 µg/ml gentamicin sulphate. The semen is centrifuged twice at 300 x g for 5 minutes then the motile and imotile sperm cells are separated on a discontinuous density gradient Percoll column by centrifugation at 1000 x g for 30 minutes.^{(35, 36)}

In vitro fertilization and embryo development

In vitro fertilization was performed successfully in the llama. Epididymal semen prepared by centrifugation in TALP medium was incubated with denuded, in vitro matured oocytes in the presence of 5 µg/ml of heparin in a fertilization medium composed of 114.0 mM NaCl, 3.2 mM KCl, 25.0 mM NaHC0₃, 0.3 NaH₂PO₄, 10 mM sodium lactate, 2.0 mM CaCl₂, 0.5 mM MgCl₂, 0.25 mM sodium pyruvate, 6mg/ml fatty acid free BSA, and 50 µg/ml gentamicin sulphate.^(34-36) A fertilization rate of 29% was obtained using this technique but only 6% and 4.7 % of fertilized eggs reached respectively the early blastocyst and hatched stage after co-culture with llama Oviductal Epithelial Cell.^{(35, 36)} There is no doubt that fertilization in vitro can be achieved in camelidae. However, it is important to develop this procedure with ejaculated semen.

Manipulation and other assisted reproduction techniques

Embryo transfer and in vitro fertilization opens new horizons to genetic improvement in camelidae. The power of these techniques can be increased several fold if embryos are manipulated before transfer. Most of the gamete or embryo manipulation procedures have for objectives the increase of the number of embryos having identical genetic make up (true twins or quadruplets or cloning), the fertilization of oocytes with sperm from males that would otherwise be infertile (sperm micro-injection), the production of embryos of a predetermined sex (sexing), or most recently, the true cloning of a superior animal.

Gamete intra-fallopian transfer (GIFT)

Intra-fallopian gamete transfer consists of placing oocytes collected from a donor into a bred or inseminated recipient's uterine tube (oviduct). Thus, fertilization and early embryo development take place in the recipient. Many oocytes may be transferred in this manner, fertilized, and then harvested at the blastocyst stage by flushing the uterus 6.5 to 7 days later.

Embryo splitting

Embryo splitting consists of cutting the embryo into two halves or four quarters to produce new embryos which have the exact same genetic make up. The embryo is split using a microblade manipulated via a micromanipulator. The embryo is fixed in one end by suction while the blade cuts through its middle. This technique has to be done with morula or early blastocyst embryos which are very difficult to obtain in camelidae, unless the embryos are flushed from the uterine tube.

Embryo cloning

Theoretically, each blastomere from an embryo is capable of giving a new complete embryo if the right development conditions are provided. Therefore, an embryo can be indefinitely reproduced with each new blastomere. This technique has been applied to cattle and sheep embryos and several offspring have been obtained. However, many problems have been reported with pregnancies using these embryos such as abnormally large calves or abnormal placentation.

Embryo sexing

Selection of the sex of the embryos before transfer is of interest to many production systems, particularly in the case of racing dromedaries. Available techniques that allow selection of the sex of the embryo can be grouped into two categories: 1) those based on the separation of X-and Y-bearing spermatozoa (sexed semen) and use of the selected population to modify the sex ratio and 2) those based on the determination of the sex of the embryos after collection, and transfer of only those that have the desired sex.

Separation of X- and Y-bearing sperm cells

The first studies on X- and Y-bearing spermatozoa were based on differences between the two populations in electrical charges, speed of motility, and mass. Most of these techniques proved very poor. A new technique for sperm sexing has been introduced recently which seems to have the most credibility. The technique, called Flow Cytometry, is based on the detection of difference in DNA content between the X and Y chromosome. DNA content is determined by the intensity of fluorescence measured by a laser beam, which is emitted by spermatozoa after staining of the DNA with a fluochrome and excitation using ultraviolet light. The difference in DNA content between X and Y sperm cells is around 3.5 to 4.5%. Sorting of cells is done by modification of the electric field which direct sperm cells to one side or the other according to the intensity of fluorescence. The machines performing this operation are called Fluorescence Activated Cell Sorters (FACS). Purity of the selected population ranges from 80 to 90%. Preliminary trials in the dromedary showed that the technique can be used to separate X and Y bearing sperm cells in this species. However, it is important to remember that because of the small number of cells sorted over a time that allows maintenance of viability, only IVF can be considered for the use of sorted semen.

Sexing of embryos

Several methods have been designed to determine the sex of embryos after collection or production in vitro. These include karyotyping of cells from an embryo or detection of substances of markers that can prove the presence of a Y chromosome or two X chromosomes. Cytogenetic determination of the sex of an embryo can be determined after culture and karyotyping of 10 to 20 cells taken by "biopsy". This technique is highly accurate but is not used currently because it takes too long and reduces the viability of the embryo.

Embryo sexing is also possible if the presence of a specific antigen found in males (H-Y antigen) can be confirmed. The test is based on an immunologic reaction directed against the H-Y antigen. The test is rapid and non-invasive but its accuracy is not very high.

The most promising method for sexing embryos remains the detection of DNA sequence using Polymerase Chain Reaction. A sequence of DNA is considerably amplified after denaturation and hybridization using a thermoresistant polymerase. The technique is very accurate and the results are obtained in 4 to 6 hours from a few embryonic cells. A commercial kit for this test is expected to reach the market soon.

Preservation of oocytes

Preservation of oocytes by freezing offers an attractive method for the conservation of genetic material from valuable females and its use several generations later for in vitro fertilization with semen from different males. Successful preservation of oocytes by freezing has been reported in the dromedary.⁽⁷⁴⁾

Cloning

No other reproductive technology has generated more interest from specialists and lay people as cloning has, both for its scientific and production potentialities as well as from the ethical questions that it raises. This is truly a self-reproduction technique which involves genetic material from any cell in the body. In simple terms, the genetic material contained in the nucleus is removed from a healthy cell and transferred into a recipient oocyte from which the genetic material (nucleus) has been removed. A special treatment is applied to the oocyte prepared in this manner in order to "jump start" division and formation of new blastomeres. Eventually a complete embryo can be transferred into the genital tract of a recipient female. The first live birth using this technique in mammals was reported in 1997 in a sheep at the Roslin Institute. The cloned sheep was derived from fetal or adult cells. The nucleus from mammary tissue cells cultured in vitro were transferred to denucleated oocytes.⁽²²⁾ Cells were induced to quiescence by serum starvation before transfer of their nuclei into enucleated recipient oocytes. Induction of quiescence in the donor cells may modify the donor chromatin structure to help nuclear reprogramming and allow development. The genetic material of the recipient egg is removed by micromanipulation before introduction of the nucleus of the donor cell and cell fusion using electric current. The electric current which is used to fuse the cells also triggers the beginning of the development of the zygote. The new embryos were then transferred to recipient sheep and resulted in genetically identical female lambs. Success rate in terms of birth of cloned animals is still too low to consider it as a production tool. Many unanswered technical and ethical questions have to be
addressed before further development can be achieved.

Transgenic animals technology

Transgenic animals refer to animals in which the DNA has been modified by introduction of a gene, or genes, from another species. Transgenic animals are created from embryos obtained by introduction of the gene of interest from one species into the host DNA. The gene is linked to a piece of DNA, called the promoter, which makes it active only in certain parts of the transgenic animal. Transgenic animals can be a valuable tool for the production of some naturally-occurring proteins that are difficult to synthesize in vitro. The protein can, for example, be secreted in the milk of transgenic animals and used for therapeutic purposes. Cloning can allow the rapid multiplication of transgenic animals making many drugs that are derived from human proteins available in sufficient quantities at a lower price.

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