Advanced Reproductive Technologies in Water Buffalo

Inherently poor reproduction and seasonality in buffalo warrants the use of advanced reproductive technologies in this species. Efforts at multiple ovulation and embryo transfer in buffaloes have resulted in the birth of calves in many countries by transfer of fresh [1-4] or cryopreserved [5] embryos, however, the efficiency in terms of superovulatory response and recovery of transferable embryos continue to be low. Attempts to improve the ovulatory responses in buffaloes have shown marginal improvements [6]. In vitro production of embryos appeared as a valid alternative to in vivo recovery of embryos. Attempts to produce embryos from follicular oocytes by in vitro maturation, in vitro fertilization and in vitro culture were successful and resulted in the birth of river [7] and crossbred (50:50 river: swamp) buffalo calves [8]. Cryopreserved in vitro produced embryos also resulted in the birth of river [9] and swamp [10] buffalo calves, including twins [11]. Embryos have also been produced in vitro by fertilization of oocytes using density gradient prepared or sex selected sperm [12-14]. Experiments on techniques such as cloning [15-18] and embryo splitting [19] have been successful in buffaloes, although the overall efficiency still needs considerable improvement. In this chapter, we describe the advanced reproductive technologies in the buffaloes except artificial insemination which has been described in a previous chapter.

1. Application and Breakthroughs

Embryo transfer techniques in water buffaloes were adopted from those in cattle with few modifications to suit the water buffalo requirements. The use and application stems from the desire to facilitate the genetic improvement program, the production of purebred animals in swamp buffalo dominated countries, and preserve endangered buffalo species like the Tamaraw (Bubalus bubalis mindoronensis). The advanced reproductive techniques that can be applied to male and female water buffaloes are oriented towards genetic improvement, conservation and transgenesis and the interplay is presented in Fig. 1 with the breakthroughs presented in Table 1.

Figure 1. Reproductive biotechnologies in water buffalo and the interplay to address genetic improvement, conservation and transgenesis.

The advanced reproductive biotechnologies in this chapter are discussed under the following topics i) in vivo production of embryos, ii) in vitro production of embryos, iii) embryo and oocytes cryopreservation, iv) intracytoplasmic sperm injection, v) sperm sexing, vi) cloning, vii) embryo sexing, viii) stem cell technology and ix) molecular genomics and nanotechnology.

2. In vivo Production of Embryos

In vivo production of embryos in buffaloes resulted in considerable interest with the first river buffalo calf being produced in the USA [1] then in India [3] and next in the Philippines [4]. Earlier attempts were reported from Thailand [20], and Bulgaria [21] with modest results due to peculiarities inherent to the buffalo [22]. Multiple ovulation and embryo transfer (MOET) optimizes the female contribution to genetic progress and it increases genetic gain by 63 to 70% per year from juvenile and adult buffalo compared to progeny testing [23]. The in vivo procedures involve induction of multiple ovulations, breeding and non-surgical recovery of embryos which are then transferred to synchronized recipients.

2.1. Single Ovulation or Induction of Multiple Ovulations

Single ovulation (SO) or induction of multiple ovulations (MO) are critical steps in the production of embryos in vivo. In SO, embryos are collected without disturbing the physiological milieu of the donor animals while MO aims to release multiple oocytes at a single estrus to optimize the reproduction potential of the donors.

In MO, the donor ovaries are stimulated to undergo extensive follicular development through the use of a hormone preparation given intramuscularly or subcutaneously to induce the growth of more large follicles than the normal number and yield a larger number of oocytes at ovulation (Fig. 2). The hormone preparations have FSH activity and studies showed that the best treatment is divided into two phases; the first phase consists of the development of follicles due to administration of FSH and the second phase consists of the rupture of mature follicles with subsequent release of oocytes, which happens under the influence of LH [3,4].

Figure 2. Superovulation protocol in water buffalo. On Day 9 to 11 after estrus, FSH (total 26 AU, Japan Denka Pharmaceutical Co., Ltd.) is administered intramuscularly at the same dose twice daily to the donors for four consecutive days at a decreasing dose. On the 3rd day, PGc is given twice intramuscularly. After observing standing estrus, first AI is done associated with intramuscular injection of LHRH-A3 (GnRH, LH, Ningbo Hormones Products Co., Ltd.). Second insemination is done 8 to 12 h from the first AI. Recovery of embryos is performed on days 5.5 to 6.5 after the first insemination [6].

Experiments on multiple ovulations in buffaloes were initiated back in 1960 [24], however, systematic studies on larger numbers of animals were conducted in the eighties [1,20]. In most of the experiments, buffaloes were superovulated using the protocols either similar to those used in cattle or used empirically. FSH, pregnant mare serum gonadotrophin (PMSG) and human menopausal gonadotrophin (hMG) are used as superovulatory hormones. In principle, any superovulatory hormone must stimulate follicular development without causing excessive ovarian edema or abnormally large follicular development and the uterine environment must be receptive to sperm transport, fertilization, cleavage and subsequent development of embryos. The ovarian response and embryo production using these hormones have been reported to be quite variable [25,26].

Water buffaloes are known to exhibit a high degree of unpredictability in their response to superovulation [4,24]. Variability in ovarian response has been related to differences in superovulatory treatments such as gonadotrophin preparation, batch and total dose, duration and timing of treatment, and the use of additional hormones in the superovulatory scheme [3,26]. Additional factors, which may be more important sources of variability, are inherent to the animal and its environment. These factors may include nutritional status, reproductive history, age, season, breed, ovarian status at the time of treatment and the effects of repeated superovulation [3,26]. Attempts to reduce the variability in ovulatory response in cattle have been modest [27] and similar results are expected in the buffalo. While considerable recent progress has been made in the field of buffalo reproductive physiology, factors inherent to the donor animal that affect superovulatory response are only partially understood. Reports on the ovulatory response and number of embryos produced with a superovulation treatment in buffaloes vary and most report a poor ovulatory response and poor embryo recovery (Table 2).

To improve the efficiency of MOET, most studies have been oriented towards developing the most appropriate treatment for induction of multiple ovulations. These trials include tests on the type, mode, dosage, time and combination of hormone administration. Results from such trials have in general shown that the ovulatory response in the buffalo is lower and less predictable than that achieved in cattle. Globally, the average number of embryos recovered per donor was 1.16 and the number of transferable embryos per donor was 0.51 [30]. Although the number of ovulations may be as high as 6.0 (Table 2), the number of transferable embryos is lower, about 2.0 per donor per flush [31-34]. One reason for such a discrepancy is the failure of ovulated oocytes to enter the oviduct [35]. Animals having 3 or more ovulations (>3CL) have been considered as responders in bovine [36]. In buffaloes, since the percentage of animals having >3CL following any gonadotrophin treatment is low, buffaloes having 2 or more than two ovulations (>2CL) are considered as responder [21,37].

Study on repeated superovulations at an average interval of 77 days over a period of 20 months produced 8.2/5.1/3.4, 6.6/3.7/2.2, 5.8/2.7/1.7, 5.1/2.4/1.7, 5.1/2.5/1.6 and 3.9/2.4/1.4 mean ovulations / total embryos / viable embryos during 1st to 6th flushings respectively indicating a drop in total/viable embryo production after first and second superovulation [38].

The poor endocrine status of water buffaloes [39], the lower number of primordial follicles on the ovary [40,41], the high incidence of silent estrus and poor palpable characteristics of the ovarian structures [42], poor fertility and seasonality of reproduction [43], the high incidence of high atresia in ovarian follicles [8] and a lower population of antral follicles at all stages of the estrous cycle [44], are some of the limiting factors which probably resulted in moderate responses of buffaloes to MOET technology [22]. Though water buffaloes are always associated with cattle, subtle differences exist and are evident by smaller size of ovaries and smaller number of follicles, poor physiological responses to superovulation treatment, less pronounced sexual behavior and sexual rhythms, and, faster development and transformation to hatched blastocysts on Day 6 to 7 [3]. The reason for early development of bubaline embryos compared to bovine embryos could partially be explained by species differences.

Follicular dynamics in the buffalo indicate differences between age groups [45], effect of management system and nutrition [46], and follicular turn-over having 2- to 3- wave-like patterns that cause variability of response to superovulation treatment [47]. Understanding the patterns of follicle development in the buffalo between age groups, season, physiological condition and management system can help improve the manipulation of the ovarian responses and enhance reproduction in this species.

2.2. Estrus Synchronization, Detection and AI

Table 3 presents the success rate of estrus synchronization procedure in some AI and ET programs. Trials to perform fixed-time AI in superovulated donors showed variability of response among animals; hence estrus detection is still an important step in any AI and ET program.

Estrus detection in water buffaloes is difficult given the high incidence of silent estrus in this animal species. Major methods to diagnose estrus include sign-based methods, manual method, bull method, chin-ball marking device, painting of tail, breeding record analysis, use of devices, or laboratory diagnosis. The use of vaginal cytology offered a low-cost, objective and practical method to predict estrus in water buffaloes [50].

Vaginal cytology.

The vaginal cytology technique is the examination of the cornification of epithelial cells in the vagina which has a direct relation to the hormonal pattern in canines. This technique was validated in estrus synchronized water buffaloes [50] where vaginal cells were collected using a 6 inch sterile swab, dehydrating the cells in concentrated methanol, fixing the cells in aceto-methanol (1:3) for few seconds, allowing the slide to dry before staining with Giemsa (KaryoMax, Life Laboratories, USA) for about 3 minutes. Initiation of AI when vaginal cell cornification was more than 40% led to successful pregnancy [50]. Figure 3 shows cornified cells in water buffaloes in comparison to other epithelial cells.

Figure 3. Images of cornified cells in water buffalo (C and D) compared to other epithelial cell stages. A. Parabasal cell dominated smear. B. Intermediate cells-dominated smear (early and late stage). C. Late intermediate (already considered cornified with small nucleus) dominated smear. D. Purely cornified cells without nucleus. Artificial insemination of two doses 12 to 18 h apart is successful when performed in both C&D conditions [50].

Sign-Based Method.

Bellowing, restlessness, tail raising when approached at the back, standing perfectly still when mounted, frequent urination, vulvar lips appear moist, red, swollen, and turgid with mucus discharge sometimes extending from vulva to feet, are all signs of estrus in buffaloes. Acceptance of the male is considered the most reliable estrus indicator. In heifer, the vulvar discharge of clear mucus in varying quantities is the most reliable single sign of estrus [51].

Manual Heat Detection Method.

Placing the palm of the hand on the rump of the animal and light massage of the vulvar lips caused the animal in estrus to react. On transrectal palpation the uterine horns are turgid and coiled and have marked tone during estrus; they are flaccid with lack of tone during diestrus.

Heat Detection by Bull.

 A vasectomized bull is placed in a pen and females in estrus migrate towards the bull. The presence of a bull in the vicinity of the females stimulates estrus behavior and is particularly helpful in detecting silent heat.

Chin Ball Marking Device Method.

A vasectomized bull [52] or an androgenized female [53] is fitted with a "chin ball marker". When the animal presses down with its chin on the back or rump region of the animal it is mounting, a spring loaded valve in the device is opened and marking fluid is released marking the back of those buffaloes that have been mounted identifying them as in estrus.

Painting of Tail Method.

 A strip of paint is well-placed on the tail head so that when another animal mounts, the paint is rubbed off marking the animals that may be in estrus. Buffaloes are required to be examined to check the paint at least once a day, otherwise heats may be missed.

Breeding Record Analysis.

This involves checking the last date of estrus in an individual buffalo using the breeding or reproduction record and calculating the time to the new expected estrus. Physiological (health and nutrition) and environmental factors, however, can cause variability in the subsequent estrus schedule.

Use of Devices.

Several devices can be used to detect an animal in estrus. Electrical resistance meter [54] is used to determine the vaginal electrical resistance. A buffalo in estrus shows the least resistance at 32.68 ± 0.46 Ohms [54]. Pedometers are used to identify movement and activity of animals in estrus. A buffalo in estrus will show an increase of up to 40% in overall movements (such as walking, running, etc.) and other activities [55]. Closed circuit television can be used to monitor the buffaloes in sheds to detect the sexually active buffalo. Sub-dermal insertion of sophisticated microchips to detect the changes in impedance, movement, activity, and temperature is also used. A rise in vaginal temperature (about 0.5 to 0.8°C) during estrus is the main characteristic feature [55].

Laboratory Diagnosis.

Various laboratory tests are available but application and use is not widespread. Progesterone assay from blood or milk samples using ELISA or RIA test are used to predict the perfect timing of estrus. Milk progesterone level is lower during the estrus period. If the decreased level persists for 2-3 days, the animal is in estrus. Vaginal smears indicate an increase in cornified acidophilic cells during the estrus period and the viscosity of vaginal mucus is lowered during estrus, thus, vaginal cytology [50] and arborization pattern of the cervical mucus can be tested simultaneously. Other methods like the use of the probes to measure conductivity [54] and endometrial biopsy can be used but the invasiveness of biopsies limit the use of this approach. Lowest electrical conductivity of the fluid in the vagina at the time of estrus [54] is the main reason for using the probe while endometrial biopsy shows that before estrus starts, phosphate activity reaches its peak and an increased level persists even 1-2 hours after the onset of estrus.

Any or a combination of the tests mentioned above can be used as aid tools in the detection of animals in estrus. Once a buffalo has been confirmed in estrus, it is inseminated.

2.3 Embryo Collection and Evaluation

The rate of development of water buffalo embryos is faster than that of cattle embryos and development to the morula stage and hatching from the zona pellucida occurs one day earlier in buffaloes than in cattle [56,57]. The resulting embryos are collected on Day 5 to 6 [3,6]. Collection on later days would result in a low embryo recovery and a high incidence of embryos at the hatched blastocyst stage.

The intermittent gravity flow method has been used to collect embryos [3,4].The donor is given an epidural anesthesia. Then a 2 or 3-way Foley catheter is inserted through the cervix into the uterine horn lumen and the balloon is inflated (Fig. 4). The catheter is then gently pulled back so that the balloon is seated; additional air is added to the balloon to completely seal the uterine horn. A Y-connector with inflow and outflow tubes is attached to the catheter. In the absence of a knob regulator, a pair of forceps is attached to each tube to regulate the flow of flushing fluid. The most commonly used medium for non-surgical embryo flushing is Dulbecco's phosphate buffered saline (PBS). The uterine horn is filled with flushing medium just enough to fill the horns (200-500 mL). The fluid in the uterine horn is agitated trans-rectally to loosen and allow the embryos to float. The flushing medium is drawn by gravity or by gentle movements [2,3] towards the collecting Encom filter. The fluid is sequentially added and removed; about one liter of fluid is used per donor. Many operators use a smaller volume and flush one uterine horn at a time. Each uterine horn is filled and emptied five to ten times with flushing medium. The volume used is according to the size of the horn. The embryos are flushed out with this fluid into an Encom filter with a pore size of 60 to 70 microns or into a large collection bottle. After flushing, the embryos in large collection bottles are allowed to settle down to the bottom of the bottle for about 30 minutes, and then an aliquot from the bottom of the bottle is collected and passed through an Encom filter. A small amount of fluid remains in the filter along with the embryos. This fluid is poured in small pre-sterilized dishes. Embryos are allowed to settle and can be searched using a stereomicroscope.

Figure 4. In vivo collection of embryos in water buffalo. Rectal canal is cleaned. A Foley catheter is inserted to the vagina and guided through the cervix up to the uterine horn. Air is introduced into the air inlet using a syringe to inflate the balloon. Flushing medium is introduced into the inlet enough to fill the uterine horn. The uterine horn is then massaged gently to allow the embryos to float then the flushing medium is drawn to the outlet to collect the embryos. (Illustration drawing by Perry Irish Hufana Duran).

Buffalo embryos can be collected at various developmental stages (Fig. 5) but viable embryos for embryo transfer are embryos in the following stages; Morula – the blastomeres closely connected and difficult to discern with the cellular mass of the embryo occupying most of the perivitelline space, the embryo at this stage looks like a cauliflower. Compact morula (tight morula) - the blastomeres are tightly connected, compact, decreased in sizes compared to a morula and cannot be discerned and the cellular mass occupies 60–70 percent of the perivitelline space. Early blastocyst - formation of blastocoel (cavity), a tiny clear space which contains fluid. The embryo increases in size and the cellular mass occupies 70–80 percent of the perivitelline space. Blastocyst - the size of the embryo has increased with the cellular mass having a prominent blastocoel that comprises more than 70 percent of the volume of the embryo. Two groups of cells are present and clearly recognizable; trophoblastic cells and the darker inner cell mass. Expanding or expanded blastocyst - the size of the embryo is increased, with no perivitelline space between the layer of trophoblastic cells and the inside of the zona pellucida. The zona pellucida becomes thinner as the blastocyst expands. A small, well compacted inner cell mass is positioned on one side of the embryo. The layer of trophectoderm cells is distinctly observed. The colour of the embryo is pale to clear because of the large amount of fluid present inside. Hatching/hatched blastocyst - the cellular mass escapes from the disrupted zona pellucida brought about by the continuous increase in size of the embryo that ultimately expands to the point of rupture of the zona pellucida. Hatched blastocysts may be spherical with a well-defined blastocele or they may be collapsed, resembling debris. Fig. 6 shows Day 6 embryos collected by the in vivo flushing method at morula to blastocyst stages while Fig. 7 shows the gross morphological appearance of an embryo.

Figure 5. Schematic presentation of various developmental stages of water buffalo embryos. (Illustration drawing by Perry Irish Hufana Duran).

Figure 6. Water buffalo embryos at Day 6 of in vivo collection. From left to right: two compact morula, morula, early blastocyst and blastocyst stages.

Figure 7. Schematic presentation of water buffalo embryo gross morphological appearance. (Illustration drawing by Perry Irish Hufana Duran).

Water buffalo embryos are morphologically similar to bovine embryos. The overall diameter has been estimated to be 150–180 μ including a zona pellucida thickness of 12–15 μ [56,57]. Not all embryos that develop to morula, early blastocyst, blastocyst and expanded blastocyst stages can be cryopreserved or transferred. Embryos are graded and classified to determine which embryos are suitable for cryopreservation (Rank A embryos) and which are best suited for immediate embryo transfer (Rank B embryos) (Fig. 8). Rank A are excellent embryos, have no visible imperfections with consistent cellular mass. Rank B embryos have a few recognizable imperfections such as few extruded cells, poor consistency or variation in the blastomere size. Embryos that show more disruption, such as a small embryonic mass with irregular shape, a large numbers of extruded or dead cells or their development appears to be delayed by one or two days with signs of cellular degeneration and disintegrated cytoplasm, are discarded and not used for ET or cryopreservation.

Figure 8 Embryo quality evaluation. (Illustration drawing by Perry Irish Hufana Duran).

Quality evaluation of embryos is based on several morphological features which is subjective. An embryo that is spherical and composed of cells (blastomeres) surrounded by a translucent gelatin-like shell known as the zona pellucida is a good landmark for embryo identification. The important criteria in evaluating the quality of embryos are: (1) shape of the embryo and the blastomeres; (2) presence of a zona pellucida; (3) size; (4) color; (5) consistency of the blastomeres, (6) stage of development of the embryo, and (7) knowledge of the age of the embryo. Water buffalo embryos are one-day ahead in their development compared to cattle [8] therefore it is suggested to collect embryos on Day 6 post insemination.

2.4 Recipient Selection and Estrus Synchronization

Recipient selection and estrus synchronization are equally important considerations in a successful embryo transfer program. The estrus synchronization between the recipient and the donor is critical to ensure pregnancy. Recipient synchronization in water buffalo is performed by IM injections of 25 mg PGF2-alpha or 0.5 mg PG-analogue administered to normal cycling females with a palpable corpus luteum. Treated animals are expected to come into estrus in two to four days with a peak on the third day. Alternatively, all recipients can be injected with prostaglandins regardless of the presence or absence of a corpus luteum then a second injection is given 11 days later, and again estrus will peak on the third day after the second injection.

The effectiveness of synchronization with prostaglandins ranges from 60 to 85 percent. Recent studies showed that the ovsynch protocol with GnRH+PGF2-alpha+GnRH is a better treatment for estrus synchronization and ovulation in buffaloes. The purpose of GnRH administration is to induce ovulation; reports showed 60–86% response among treated buffaloes [59,60] and the interval between GnRH administration and ovulation is 33±8.3 h. The average percentage of synchronized estrus obtained was 91.7%, and the pregnancy rate following embryo transfer was 30.4% [58]. GnRH also enables the early corpus luteum to develop, mature and enhance LH levels [59].

Before applying prostaglandin to buffaloes to regress a functional CL, one can verify plasma progesterone concentrations, view the CL through ultrasonography to assess its size and confirm the ovarian status of the animal [61]. If the CL is already regressing, prostaglandin treatment will not accelerate its demise.

Prostaglandin treatments are administered either IM or SC; administration by intra vulvo-submucosal injection proved equally satisfactory, even at reduced dosages [62]. An inadequate luteal phase may occur due to a short lifespan of the CL. This is characterized by insufficient progesterone production which in turn may explain the unsatisfactory response to prostaglandin administration for control of reproductive function [63].

Body condition score is an important consideration in the selection of recipients. A large-framed, healthy, reproductively mature young buffalo with a minimum of two normal cycles is ideal. Animals in poor body condition and early post-partum animals do not make suitable recipients. Recipients should preferably be gaining 0.1–0.2 kg per day and if possible, animals should be vaccinated against the known abortion-producing diseases prevalent in their area.

2.5 Embryo Transfer

Embryo transfer involves the use of an embryo transfer gun with inner and outer sheaths to ensure hygiene of both the embryos and the recipient. The recipient is selected based on estrus; preferably the selected animal should have exhibited estrus on the same day or just before the donor. To ensure a high success rate, only healthy embryos judged according to their color, size, and consistency are to be transferred.

Before carrying out the embryo transfer, the presence of CL on the ovaries must be confirmed either by trans-rectal palpation or by transrectal ultrasonography. To ensure the easy penetration of the ET gun into the cervix, a cervix expander is first inserted into the vagina of the recipient animal [9]. After expanding the cervix, the expander is removed and the ET gun is inserted into the entrance of the cervix with an outer sheath to ensure that the inner sheath is clean when it is passed through the vulva and the vagina. The tip of the outer sheath is punctured and the gun with the inner sheath is inserted into the cervix and pushed gently until it reaches the uterine horn ipsilateral to the ovary bearing the CL. For the deposition of the embryo, the tip of the gun is inserted up to 5–10 cm beyond the external bifurcation so the embryo is deposited into the uterine horn (Fig. 9). A hand inserted trans-rectally guides the ET gun into the right position in the uterine lumen and holds the gun in place while the embryo is being released.

Figure 9. Schematic presentation of embryo transfer in water buffalo. (Illustration drawing by Perry Irish Hufana Duran).

3. In Vitro Production of Embryos

In vitro production is the best alternative for the production of embryos for ET and to use in research applications. This technique can be used to test different advanced technologies such as cryopreservation, intracytoplasmic sperm injection (ICSI), somatic cell nuclear transfer (SCNT), IVF with sexed-sperm, embryo splitting, screening of genetic defects as well as genetic manipulation, basic research in developmental physiology, and exploitation for emerging biotechnologies such as transgenesis and cloning.

The in vitro embryo production (IVEP) involves a series of integrated sequential steps: 1) collection of oocytes by retrieval from abattoir derived ovaries or by ovum pick up from live donors, 2) selection of developmentally competent oocytes and in vitro maturation (IVM) of the selected oocytes, 3) sperm capacitation and in vitro fertilization (IVF), and 4) in vitro culture for embryo development (IVC). In addition, other technologies such as ICSI with sex-sorted sperm cells and SCNT [15] can be carried out.

Events and processes of IVEP must be done in aseptic conditions to avoid contamination. A laminar flow cabinet is appropriate. All materials and media that come into contact with gametes and embryos should be sterile to avoid bacterial and fungal contaminations. In the absence of sterile, disposable supplies and materials, autoclaving at 121°C for 15 minutes is appropriate both for plastic and glassware wrapped in aluminum foil or autoclavable plastic bags. For culture media, sterilization by filtration using syringe filters with a 0.2 micron membrane is sufficient. To minimize stress to the gametes, in vitro manipulation procedures are done at appropriate temperature (35 to 37°C), pH (7.1 to 7.4), osmolarity (280 to 300 mOsmol), and minimum exposure to UV light. In vitro culture is done in a humidified CO2 incubator at 38-39°C [22]. The sequential steps for in vitro embryo production (IVP) are described below.

3.1 Ovary Collection and Oocyte Aspiration

Unless the slaughtered animal has a known genetic value confirmed by its record, the collection of oocytes from abattoir derived ovaries does not guarantee the acquisition of the best genetic material. However, careful examination and evaluation of the records of the ovary donors, if available, can be used as reference. Oocytes from unknown sources are used for research and educational purposes only.

Ovary collection requires proper handling to preserve the viability of the oocytes. The common storage medium used is physiological saline: 0.9% sodium chloride in distilled water with 100 μg streptomycin/mL and 100 IU penicillin/mL [9] or without antibiotics [64]. The temperature of the storage medium depends upon the length of storage of the ovaries; body temperature is appropriate if oocyte collection will be done immediately, higher temperatures (>39°C) are detrimental. If oocyte aspiration will be carried out one hour after ovary collection but no more than six hours, 25 to 33°C is best [57]. For longer periods of time, 15°C is the preferred temperature for ovary storage [65]. Lower temperature preserves the viability of the ovarian tissue for a longer period of time. Upon arrival in the laboratory, all ovarian manipulations must be done at 37°C to maintain viability of oocytes. Washing of the ovaries using fresh physiological saline upon arrival to the laboratory is necessary to ensure that the ovaries are clean prior to aspiration of the oocytes.

Oocyte recovery can be done by aspiration (using a disposable sterile syringe), ovarian slicing, or follicle puncture. Slicing yields a larger number of collected oocytes [44,66,67], however, this method is not convenient because of the high percentages of fair, poor and denuded oocytes obtained and the high incidence of contamination. Follicular aspiration is considered the best method. The mean number of good oocytes collected from a buffalo ovary ranges from 0.43 to 3.3 oocytes/ovary [44,57,66,68-70] and the variability is dependent on the competence and efficiency of the person doing the aspiration, and also the breed, health of the donor, size of the ovary, the number of follicles accessible on the ovary, presence or absence of a CL and the season of the year [22]. The total number of oocytes/culturable grade oocytes recovered from buffalo ovaries is lower than in cattle at any given time. This limitation is due to the inherent low follicular reserve in buffaloes. It has been shown that buffalo ovaries have a smaller population of recruitable follicles with an average of 12,000 primary follicles [40,45].

The quality of oocytes is important. The aspiration process can mechanically damage the oocyte in two ways: 1) the size of the aspiration needle, and 2) the aspiration vacuum used to recover the oocytes. A sterile 18-gauge needle attached to a sterile 10 mL plastic syringe pre-loaded with oocyte holding medium is used. To collect and hold oocytes, a sterile tip with bore opening bigger than 150 μ (i.e., bigger than the size of an oocyte with its surrounding cumulus cells) must be used to ensure collection of the oocytes without damaging the surrounding cumulus and granulosa cells. The cumulus cells play an important role in the developmental competence of the oocyte. To avoid mechanical damage of the surrounding cumulus cells, it is necessary to apply low pressure aspiration.

The oocyte holding and washing mediums used by various authors are: 1) Tissue Culture Medium-199 (TCM199) with 10% fetal calf serum (FCS), buffered with 15 mM HEPES and 5 mM sodium bicarbonate [71]; 2) Pre-warmed modified phosphate buffered saline (m-PBS) with 3 mg/mL bovine serum albumin [11,72], or 5% (v/v) heat inactivated FCS [73]; 3) HEPES-buffered modified Tyrode’s medium [74] supplemented with 3 mg/mL bovine serum albumin, 0.2 mM sodium pyruvate (Sigma) and 50 μg/mL gentamicin sulfate (Sigma) [75].

BSA and serum are provided to avoid adherence of the oocytes to the surface of the container and to provide energy substrates. Follicular aspirates are dispensed in sterile conical tubes to allow the oocytes to settle down onto the bottom of the tube to facilitate retrieval after the aspiration. Oocytes are then washed by sequential transfers into three small sterile culture dishes (35-90 mm) containing 2.5 mL washing medium. Prior to the final IVM droplet, they are also washed in by sequential transfers (3-5 times) into small microdrops (50-100 μL) of IVM medium in the sterile culture dishes [57,76].

Good quality oocytes are recovered from ovaries of cyclic buffaloes at days 1-3 and 10-16 of the estrous cycle [68], in the presence of a CL [69,77], and in animals with a body condition score of 3-5 [78].

3.2 Ovum Pick Up (OPU)

OPU is carried out on live donors. The technique involves ultrasound-guided follicle aspiration for recovery of oocytes. This technique allows for the repeated pick-up of immature oocytes directly from the ovary without any major impact on the female donor. The use of these oocytes in IVM/IVF programs allows a greater use of genetically valuable females.

OPU is an extremely versatile technique because it can be applied to donors of all ages from two month-old calves to very old cows with exception of pregnant animals after the third or fourth month of pregnancy, animals with severe ovarian hypoplasia or during the immediate postpartum period before ovarian activity is restored [79]. OPU appears to be the most reliable source of genetic material to enhance female contribution to genetic progress.

To collect oocytes by OPU, a 51 cm long 0.1 cm diameter stainless steel needle with ultrasound echo tip is aligned to a vaginal ultrasound probe (Fig. 10) to aspirate the oocytes from the follicles. The needle is attached to a 50 ml collecting tube by silicon tubing anchored to a vacuum pump (e.g., Fujihira Co. Ltd., Japan) set to produce a negative pressure of 60 mmHg during aspiration. Prior to aspiration, the collecting needle and tubing are pre-loaded with the collecting medium or modified Dulbecco’s Phosphate Buffered Saline (mPBS) with 3% Bovine Serum Albumin (BSA), 50 μg/mL gentamicin and 20 μg/mL heparin. In the absence of BSA, 1% Polyvinyl Alcohol (PVA) can be used.

The technique of OPU utilized in buffaloes has been the same described for cows [80]. Briefly after epidural anesthesia and evacuation of the contents of the rectum [81], the operator places the probe in the vagina covered with a sleeve and places it lateral to the cervix. The other arm is placed in the rectum and the operator brings the ovary closer to the probe to visualize the ovarian follicles. Once the follicles are visualized, the long needle is penetrated through the vaginal wall to bring the needle close to the visible follicles. The follicle wall is carefully punctured with the needle and the follicular fluid is aspirated through the vacuum pump attached to the needle. The needle can then be withdrawn and the procedure can be repeated on all follicles and then move over to the other ovary. OPU involves the use of the appropriate size needle and aspiration by hitting only the base of the follicles avoiding too much penetration of the needle into the ovarian medulla.

Figure 10. A transvaginal ultrasound guided equipment (vaginal probe) with the aspiration needle.

The follicular aspirate is maintained at 38°C in a warming block and transported to the laboratory in the same condition. Cumulus-oocyte complexes (COCs) are retrieved by putting the aspirate through an EnCom filter discarding the aspiration medium and washing the follicular aspirates with a pre-warmed mPBS with 3% BSA or 1% PVA without heparin.

Different schedules can be used for oocyte recovery allowing the system to adapt to different practical situations. A twice-a-week collection allows for the maximum recovery of oocytes of suitable quality for embryo production in a given time interval [82]. A once-a-week collection results in the recovery of a smaller number of oocytes (of lower quality) that have already undergone cumulus expansion and atresia [83]. This is because in the twice-a-week collection, if all visible follicles are aspirated, no dominant follicle develops. In most once-a-week collections, a dominant follicle is present at each successive collection causing the regression and degeneration of the subordinate follicles. Also, the donors can come into estrus while this is never the case in the twice-a-week schedule. Some authors prefer to combine superovulation with OPU and to recover the oocytes before the onset of estrus [79] but the procedure can be repeated at best every two weeks. The average yield of embryos can be higher per OPU but the total number over a period of time is lower than for the twice-a-week collection. Prior stimulation in buffaloes with gonadotrophins [84,85] or bovine somatotropin [86,87] or pharmacologically synchronized follicular waves [88] before OPU are known to increase the number of medium and large sized follicles, however, the embryo production after IVEP was not affected. The OPU-IVEP during the breeding season yielded a better oocyte recovery and the resultant embryos are of better quality and higher development potential [89,90]. OPU can have its own therapeutic effect on infertile donors, especially those affected by ovarian cysts [83]. Virtually all non-pregnant donors return to estrus within two weeks from the last oocyte collection and can be inseminated successfully.

OPU has no side effects even after twice-a-week collections for over a year [81]. In some cases though, a minor hardening of the surface of the ovaries can occur after several months of repeated collections. However, in one study there was a decline in the follicle recruitment and oocytes collection after the first 2 months of recovery [91].

Production of offspring combining OPU with IVEP and vitrification for direct ET appears a promising combination of good applicability in breeding and production of calves [92]. Studies in Italy and China noted the potential impact of OPU technique combined with IVEP in the diffusion of embryo transfer procedures in the field and on genetic improvement of buffalos [93,94]. In spite of the current application of OPU, it seems clear that more efforts have to be made to optimize both repeated OPU retrieval and, particularly, the current IVM procedures, which currently appear to be the major limiting factor for a satisfactory IVEP.

3.3 Oocyte Selection and In Vitro Maturation

Oocyte selection and in vitro maturation are the foundations of in vitro embryo production. The quality of oocytes, the pH, and osmolality of the culture medium and the temperature and gas phase of the culture environment are important factors [57]. One has to bear in mind that fully grown, prophase I-arrested oocytes collected from the ovaries are maintained in meiotic arrest within ovarian follicles by inhibitory factors produced by the follicle. When oocytes are removed from Graafian follicles and cultured in vitro under suitable conditions, they resume meiosis spontaneously and progress to the metaphase II (MII) stage (Fig. 11). The ability of buffalo oocytes matured in vitro to develop to the morula/blastocyst stage is lower compared to in vivo-produced embryos. The decrease in the developmental competence of in vitro-matured oocytes may be due to insufficient cytoplasmic maturity brought about by the limitations in the in vitro culture environment. Hence, continuous efforts are being made to improve the current culture systems and enhance the developmental competence of buffalo oocytes and embryos in vitro.

In selecting oocytes for IVM, pre-warmed holding and washing medium must provide the necessary nutrients and environment needed by the oocytes to sustain their developmental competence. Various media used were reviewed [79,93,95]. Oocytes are selected by the compaction of the cumulus-corona investment and homogeneity of the ooplasm (Fig. 12). The presence and quality of cumulus cells surrounding the oocytes is essential to facilitate the transport of nutrients and signals into and out of the oocytes. Oocytes with a compact cumulus cell mass (Fig. 13a) require a longer (24 to 26 h) period of IVM and those with loose cumulus (Fig. 13b) require a shorter (20 to 22 h) period of IVM for optimum blastocyst development [57,96]. The granulation of the ooplasm (Fig. 14a and Fig. 14b) has no significant difference on the developmental competence of the oocyte.

Figure 11. Nuclear status and classification of water buffalo oocytes; A) Germinal vesicle (GV), nucleus is enclosed by a membrane. B) Germinal vesicle breakdown (GVBD), condensation of chromosomes and disappearance of the membrane. C) Metaphase I (M-I), spindle formation. D) Anaphase I (A-I), separation of homologous chromosomes. E) Telophase I (T-I), extrusion of the first polar body. F) Metaphase II (M-II), polar body in the perivitelline space. (Hufana-Duran, 2008. Ph.D. Dissertation, University of Tsukuba, Japan).

Figure 12. Quality of water buffalo oocytes classified based on the morphology of the surrounding cumulus cells; A) Rank A – surrounded by ≥5 layers of dense cumulus mass. B) Rank B – surrounded by 2-<5 layers of dense cumulus mass. C) Rank C – surrounded by <2 layers of irregular cumulus mass. D) Rank D – denuded or free from cumulus cells. E) Rank E – surrounded by expanded cumulus mass [57].

Figure 13. Compactness of the surrounding cumulus cells. A) Oocyte classified as with compact cumulus cells, and B) oocytes with loosened cumulus cells. [57].

Figure 14. Granulation of ooplasm. Oocytes classified as with A) homogeneous or evenly granulated ooplasm, B) heterogeneous or unevenly granulated ooplasm. Note the darker portion on the side of the ooplasm in oocytes in B plate [57].

Oocyte diameter and the size of the donor follicle are positive indicators of oocytes’ developmental competence; oocytes with a diameter <100 μm lack developmental competence and most likely fails to develop to MII after IVM [57]. Oocytes with a diameter ≥100 μm developed to MII, cleaved after IVF and those with a diameter ≥120 μm have the highest chances to develop to the blastocyst stage. The size of the donor follicle is linearly correlated with oocyte developmental competence with follicles ≥6 mm containing highly developmentally competent oocytes. The Brilliant Cresyl Blue (BCB), an indicator for glucose-6-phosphate dehydrogenase activity, can be used to identify oocytes that are still at the growing phase [57]. Growing stage means the oocyte is at the stage of accumulating the developmental competence to undergo meiosis. Studies [57,97] reported that oocytes with highly compact cumulus cells (Fig. 13a and Fig. 14a) showed significantly higher BCB negative (39.1%) compared to the oocytes with loose cumuli (Fig. 13b) (8.6%) indicating that some oocytes with very compact cumulus cells are still at the growing phase of development, thus, requiring a longer period of oocyte maturation [96].

A major factor affecting in vitro mammalian embryo development is increased oxidative stress [98]. An ultrastructural study of buffalo oocytes showed an abundance of cytoplasmic granules characterized by significant lipid content [57,99] that imparts the cytoplasm of oocytes and embryos a dark appearance at light microscopy and probably renders the buffalo oocytes and embryos more sensitive to oxidative damage.

Handling of the oocytes requires the use of sterile glass pipettes either mouth-controlled or hand-controlled. Micro-pipettes and embryo exhausters can also be used [76]. The holding pipette is washed with pre-warmed holding medium at least three times prior to holding the oocytes to avoid contamination. Selected oocytes are held in a holding medium and washed at least three times (to remove the debris) prior to final placement in medium for IVM. The use of glass pipettes allows preparation of the desired bore diameter that suits the size of the oocytes during different applications i.e. 180 to 200 μm for IVM, 150 to 180 μm for IVF, and 110 to 120 μm when desired to denude the oocytes.

The culture medium has marked effects on the fertilization rate, development of pre-implantation stage embryos, resistance to cryopreservation stress, and on full-term development after embryo transfer. A variety of media are available for use in the culture of buffalo oocytes and these could be categorized into simple and complex [93]. Simple media are usually bicarbonate-buffered containing physiological saline with pyruvate, lactate and glucose, and they differ in their ion concentration and in the concentrations of the energy sources [100]. Complex media contains the basic components of simple media with the addition of amino acids, vitamins and purines. Chemically defined media have known chemical components entirely free of animal-derived components including serum and represent the purest and most consistent cell culture environment suitable for the in vitro cell culture [93]. The critical factor in the in vitro culture environment is the provisions of the support needed that signals and enhances the mechanisms to acquire developmental competence by the oocyte. TCM 199, Ham’s F-10, CR1aa, and CR2aa, MEM, mSOF, and RPMI-1640 are used as basic media and are supplemented with either serum [100], hormones [72,101], growth factors [76,102,103], antioxidants, follicular fluid [64,104] and a controlled level of antibiotics to provide protection from bacterial contamination. Several studies on the best culture system have been carried out with promising results, and resulting in the birth of live calves [9,13-16,105]. For maximum oocyte in vitro maturation, TCM 199 supplemented with serum, FSH, and cysteamine is optimum in producing maturation and subsequent embryonic development after insemination [106]. Table 4 presents examples of media used for in vitro maturation of buffalo oocytes with corresponding blastocysts development rate.

Serum has been used as an effective supplement in the in vitro culture systems of buffalo oocytes and even embryos. Animal serum or albumin is routinely added to culture media as a source of many nutrients required by cells to proliferate and survive. Among other things, serum contains proteins (i.e. albumin) which serve as carriers and protective agents for other molecules by binding with vitamins such as pyridoxal (active form of vitamin B6 that serves as cofactor in amino acid metabolism), fatty acids such as oleic, linoleic, linolenic, arachidonic, myristic and palmitic acids, and ions such as copper [109]. It increases the solubility of these molecules, maintains them in chemical states that are unstable in an unbound form, and protects them from oxidation. Serum also promotes cumulus cell-oocyte uncoupling by retaining the hyaluronic acid within the COCs in a manner that results in cumulus mucification. This uncoupling could be responsible for stopping the transfer of oocyte maturation inhibition factor via gap junction [110]. However, the use of serum supplementation poses some technical disadvantage because of its undefined nature, batch-to-batch variability in composition, and the risk of contamination. This has led to the use of chemically defined or serum-free media formulations [111,112]. Nonetheless, buffalo oocytes can be matured in TCM-199 medium supplemented with one of the following sera at 5 to 10% concentration: (1) buffalo estrus serum (BES), (2) superovulated buffalo serum (SBS), (3) fetal bovine serum (FBS) and (4) steer serum (SS) but cleavage and blastocyst rates are higher (P<0.05) in SBS [101] as reviewed earlier [95,106].

The culture environment inside the incubator is important in ensuring a smooth maturation of oocytes. A humidified environment at 38.5°C and 5% CO2 with relative humidity of 95% is optimum for efficient in vitro culture. Temperature of 36.5°C and 39.5°C could significantly decrease the maturation rate of oocytes [113].

Efficiency of the culture medium on the maturation of oocytes is measured by visual observation under stereo microscope using the following parameters; 1) the expansion of the cumulus cells, and 2) the formation of the first polar body. The presence of the first polar body could be examined by partly denuding the oocytes from the surrounding cumulus cells. These methods are helpful if the oocytes are to be used for IVF. Depending on the purpose, maturation rate of the oocytes could be examined by checking the nuclear maturation of the oocytes by fixation using 1:3 aceto-ethanol, dehydration with absolute ethanol once oocytes are laid under a cover slip, staining with 1% aceto-orcein and de-stained with aceto-glycerol (glycerol: acetic acid: distilled water = 1:1:3 v/v) [57].

3.4 Sperm Capacitation and In vitro Fertilization

Sperm capacitation is the physiological changes spermatozoa must undergo in order to have the ability to penetrate and fertilize an egg. This event consists in unmasking the receptors on the sperm itself, rendering it capable of reacting to the environment of the female genital tract [114]. Capacitation involves removal of the glycoprotein coat to uncover receptors that may recognize the chemicals in the female reproductive tract that in turn initiate changes in both the motility (in the form of hyperactivation) as well as the morphological changes involved in the acrosome reaction [115]. In vitro, it is necessary to provide the capacitation signal to the sperm cells in order to produce fertilization. The medium employed in IVF systems must be capable of providing the secondary oocyte and the capacitated sperm with the conditions, which will permit sperm penetration to occur readily. It was reported that 2X detoxified uterine fluid in Riggers Whitten Whittingham (BWW) medium; pH 7.4 at 37°C induces sperm capacitation [116]. There are different media described that could enhance sperm capacitation in vitro such as Fert-TALP [73,117] but Brackett & Oliphant medium was found marginally better [69,118] with sperm motility enhancers such as 2.5 mM caffeine and 10 μg/mL heparin [7,9,69,119,120,] or theophylline [57] or a mixture of phenylephrine, hypotaurine and epinephrine [76].

Depending on the quality of semen, the presence of numerous dead sperm in the IVF environment is not beneficial as the dead sperm secrete factors detrimental to the live sperm and oocytes. To separate the live from the dead sperm population, swim up procedures can be used [117], however, ion-exchange filtration was found to be better [121]. Discontinuous density gradients of percoll [76] or silica particles [12] can also be used.

Sperm concentration is an important factor for in vitro fertilization. Too many sperm could result in polyspermy while too few sperm could result in low fertilization rate. IVF in buffaloes is carried out at 1 to 2x106 sperm cells/mL. Adjustment of the sperm concentration can be done by determining the sperm concentration of the frozen-thawed-washed sperm cells using a spectrophotometer or counting the sperm cells using a haemocytometer or Neubauer chamber [22,95].

Usually, IVF droplets are diluted 1:1 v/v with sperm dilution solution to form an IVF medium with final sperm concentration of 1x106sperm cells/mL, 5 mM caffeine, 2 units heparin/mL and 5 mg BSA/mL. To execute IVF, in vitro matured oocytes are partly freed from cumulus cells to enhance sperm penetration and promote higher fertilization rate. The detached cumulus cells are retained on the IVM droplets to develop a cumulus cell monolayer for embryo-somatic cell co-culture system.

Important laboratory tests to assess sperm quality parameters include: CASA to assess sperm motility, eosin-nigrosin staining technique to assess the vitality of sperm cells, chlortetracycline (CTC) fluorescence assay and Pisum sativum agglutinin (FITCPSA) [118,122] or trypan blue/giemsa [123], staining techniques to check capacitation and acrosome reaction, and hypo-osmotic swelling tests to check the functional integrity of the plasma membrane which is also associated with sperm fertility [124] are utilized.

Sperm-oocyte co-culture for in vitro fertilization is carried out for 6 to 18 hours. The duration of in vitro fertilization is dependent upon the composition of the in vitro fertilization medium and the sperm concentration. It is necessary to examine the best duration of sperm-oocyte co-culture as differences exist depending on the IVF media formulation, sperm concentration, and the bull used [125].

3.5 In vitro Culture for Embryo Development and Grading

The culture of embryos in vitro requires the necessary nutrients and appropriate environment so that the fertilized oocytes can undergo cleavage divisions and be able to reach the blastocyst stage of development. In this respect, several culture media have been tested and all resulted in the development of blastocysts with varying degrees of success: CR1aa, CR2aa media, TCM-199, MEM, RPMI-1640, and mSOF media [93,106]. The effectiveness of each medium formulation depends mainly on provision of the appropriate combination of antioxidants, co-culture, growth factors and gas phase. The methods used in bovine for IVC of embryos by co-culture with cumulus cells [126] and the sequential media system containing pyruvate and lactate and different concentration of serum and presence of glucose [127], were adopted in water buffaloes and these resulted in full-term development after embryo transfer [9-11].

The success of IVF is assessed by the cleavage rate on the second day of in vitro embryo culture [101,104]. Cleaved embryos are separated from the uncleaved ones and are further cultured to reach blastocyst development. Depending on the culture system, embryo development is monitored daily and the culture medium renewed every 2 days or the culture is left without disturbance and development to blastocysts is checked on the 6th or 7th day of IVC [57,93]. During embryonic development, cell numbers increase geometrically (1-2-4-8-16- etc.) with blastomeres exhibiting equal sizes and consistency in good quality zygotes (Fig. 8). Synchronous cell division is generally maintained to the 16-cell stage. After that, cell division becomes asynchronous and finally individual cells possess their own cell cycle. The cells composing the embryos are termed blastomeres and are easily identified at the 16-cell stage as spherical cells. After the 32-cell stage (morula stage), embryos undergo compaction. As a result, individual cells in the embryo are difficult to discern. The most obvious morphological manifestation during compaction is the loss of a concise cellular outline. After compaction, the dynamic cell differentiation begins with the formation of the blastocyst with two distinct cell types; the inner cell mass and the trophectoderm cells, are formed and differ in their morphology, biochemistry, developmental potential and eventual expression.

It has been noted that embryo cryopreservation and the potential impact of the OPU technique combined with IVEP critically contribute to the diffusion of embryo transfer procedures in the field and on the genetic improvement of buffaloes [94].

3.6 Important Considerations in IVEP Works

The success rate in the production of embryos in vitro is affected by several factors: 1) source of oocytes (age, body, and health condition of donors), 2) quality of the oocytes (presence of cumulus cells and granulation of ooplasm), 3) component, pH, and osmolarity of the in vitro culture medium, 4) in vitro culture environment (temperature, humidity and gas) [93,106]. These factors significantly affect post-fertilization embryo development. Oocytes from juvenile donors lack the developmental requirements while those from adult donors have a high incidence of chromosome abnormalities. Thus oocytes from very young or old animals are collected only for research. Animals that are too thin or too fat or are sick will most likely provide low quality oocytes. Changes in temperature, pH, osmolarity and contaminant-factors which may affect viability must be guarded carefully. Oocytes and especially embryos do not tolerate temperatures above body temperature (39°C) very well.

Oocytes and embryos require a physiological pH ranging from 7.1 to 7.4. The holding, washing and culture media need to be within this range. Phosphate buffered saline (PBS) supplemented with serum or BSA is commonly used because under atmospheric conditions the change in pH is negligible. Bicarbonate (NaHCO3) buffered media like TCM 199, Ham's F-10, MEM, RPMI and others are also used but speed of operation is needed as extended time under atmospheric conditions high in CO2 would result in large shifts in pH (rising pH) that highly compromise the quality of the oocytes and embryos [57].

The osmolarity (the concentration of salt) of the medium also affects the viability of oocytes and embryos. If the osmolarity of the media is below that of the in vivo environment, oocytes and embryos will absorb water resulting in swelling to reach osmotic equilibrium. This could result in rupture of the cell membrane which highly compromises the developmental competence of the oocytes and embryos. Conversely, if the salt concentration is high, the oocytes and embryo will shrink in size (dehydration) causing a reduction in metabolic activity. If PBS is prepared from a powdered mixture, care should be taken to correct the amount of water added. The normal osmolarity of uterine fluid is 270–300 mOsm [128].

Exposure of embryos to ultraviolet rays should be avoided to prevent cellular death. The embryology room must provide an aseptic environment favorable for handling of oocytes, sperm cells, and embryos. Use of insecticide sprays is detrimental. In using ethylene oxide gas for sterilization of equipment, sufficient aeration time must be ensured since the residue would be detrimental to the oocytes and embryos.

3.7 Pre-antral Follicle Culture

With the limited number of oocytes retrieved from buffalo ovaries and a number of potential application on transgenesis, conservation of rare species, formation of genetic material banks and to study folliculogenesis; attempts to culture pre-antral follicles were initiated [129,130]. The preantral follicles are isolated by micro-dissection of abattoir derived ovaries and 1% trypsin digestion of ovarian cortical slices [131]. Initial studies cultured small and medium sized follicles [130], however, later studies used medium and large sized (150-500 μ) follicles for culture [131-135]. The medium commonly utilized is minimum essential medium (MEM) supplemented with steer serum (10%), FSH (5 μg/mL), sodium pyruvate (0.23 mM), glutamine (2 mM), hypoxanthine (2 mM), insulin-transferin-selenium (1%) and antibiotics [130,131,135,136]. The follicles are placed in microdrops in a dish and the culture is carried out in a CO2 incubator for 80-100 days with the medium being changed every other day [131] or every 10 days [132]. Studies showed that the growth of preantral follicles cultured in groups is better than those cultured individually [130,132]. Growth factor and hormonal supplementation in the culture medium enhanced the growth [130,137]. Addition of somatic cells such as oviductal mesenchymal cells (OMC), granulosa cells (GC) or cumulus cells (CC) supported higher growth rates [131,133,136], with maximum development and survivability being observed in CC and OMC. In one study, preantral follicles were cultured with antral follicles (AF) and this supported higher survival, growth rate and antrum formation. The effect of nitric oxide on development of buffalo preantral follicles was positive at lower concentrations [138,139]. Study on the preantral follicle culture from buffalo fetuses (5 to 9 months) [129] showed follicle growth and although the follicular architecture was preserved in nearly 17% of the follicles, antrum formation failed. Difficulty in isolation of preantral follicles, fewer numbers, poor growth in culture and poor subsequent development of retrieved oocytes are some of the problems encountered in preantral follicle culture [129,136].

4. Embryo and Oocyte Cryopreservation

Cryopreservation is a process where cells are preserved by cooling to low sub-zero temperatures allowing transportation, long-term storage, conservation, and programmed utilization. This technology is important for the cryobanking of animal’s germplasm from endangered species and exploitation of genetically superior sires through AI and embryo transfer. Cryopreservation of sperm is an old technology with large scale commercial application. Cryopreservation of embryos is an advanced technology that has achieved considerable success with birth of calves after transfer of cryopreserved embryos [9-11,17]. Cryopreservation of oocytes and somatic cells has practical application for cloning and gene banking. Even though sperm, embryo, and somatic cell cryopreservation succeeded and these are now routine activities, oocyte cryopreservation remains a challenge. Recent studies [140,141] showed improvement on development to morulae and blastocysts after IVF of oocytes cryopreserved by slow-freezing and vitrification but the efficiency remains lower than that for fresh oocytes [141]. Efficiency of oocyte cryopreservation requires further refinement before it can be used for routine application. The high lipid content of buffalo oocytes [57] may influence the success of cryopreservation [142].

There are two basic techniques currently used in the field of cryopreservation; 1) slow freezing and 2) vitrification. Cryopreservation by slow freezing is a process where extracellular water crystallizes, resulting in an osmotic gradient that draws water from the intracellular compartment until intracellular vitrification occurs. This requires slow freezing equipment programmed to undergo gradual temperature changes to sub-zero to carry out the cryopreservation process. In cryopreservation by vitrification, both intra and extracellular compartments apparently vitrify, after cellular dehydration has already occurred. The process does not require invasive equipment. The correct formulations of cryoprotectant, temperature and exposure time are the main secrets of successful cryopreservation. Owing to these differences, the terms freezing and thawing are relevant to the slow freezing process while cooling and warming are relevant to vitrification. Vitrification has been considered an efficient method in the cryopreservation of biological samples that are particularly sensitive to chilling injury like buffalo embryos [9,143] and oocytes [144,145].

Vitrification utilizes a minimum volume and higher levels of cryoprotectants to allow ultra-rapid cooling and warming to overcome chilling injury and cytotoxicity from cryoprotectants. Several approaches are used; 1) traditional French straw [146,147], 2) open-pulled pointed end straw [9,144], 3) cryotop [143,148], and 4) solid surface vitrification method and cryoloop [149]. These methods allow ultra-rapid cooling (from 25°C to sub-zero temperature) and warming (from sub-zero temperature to 25°C) on account of extremely small volume used (<0.1μL).

The vitrification method [150] modified to suit water buffalo embryos [151] and oocytes [140,141] have been shown to be successful in the cryopreservation of buffalo embryos [9-11] and oocytes [144,145]. The vitrification solution used was EFS40 (ethylene glycol, 40%, v/v; ficoll, 18%, w/v; sucrose, 0.3 M) and the procedure was carried out in a working room at temperature of 25°C. All solutions used were equilibrated at the same temperature for at least 3 h to avoid exposure of the embryos to temperature fluctuation. To facilitate the vitrification process, an open pulled French straw with one end sharpened to have a pointed tip [9] is prepared to load the embryo. To vitrify, the embryos are washed with Dulbecco’s phosphate-buffered saline containing 5.56 mM glucose, 0.33 mM pyruvate, 100 units penicillin/mL and 3 mg BSA fatty acid free/mL (referred as PB1 medium), equilibrated in 10% ethylene glycol in PB1 medium for 5 minutes then exposed to EFS40 and placed on the tip of the pointed-shaped open straw within a period of 30 seconds and rapidly plunged in liquid nitrogen [10]. Straws are kept in a liquid nitrogen tank until a recipient is ready for embryo transfer. For higher success rate, only Rank A embryos (embryos with intact zona, no visible imperfections and consistent cellular mass) are considered for cryopreservation and Rank B embryos (embryos with few imperfections) are best for transfer without cryopreservation. For ET, embryo straws are recovered from liquid nitrogen tanks and the pointed end where embryos are loaded is directly warmed in 0.5 M sucrose solution in PB1 medium at 25°C for re-expansion within 5 min. Re-expanded embryos are washed in PB1 medium and loaded in 0.25 ml French straw for transfer. Procedures for vitrification of oocytes have been mentioned [148,152] with 8M concentration of ethylene glycol being suggested as the optimum cryoprotectant [144,145]. The presence of cumulus cells with the oocytes provides additional protection from cryoinjury [153].

Cryopreservation of embryos by slow-freezing is a proven procedure [5] but adoption has not prospered because of the equipment needed and the time-consuming approach. Conversely vitrification is easy to perform and embryos can be easily thawed and transferred directly [9].

Cryopreservation also plays an important role in the preservation of ovarian tissue [154]. The cryopreservation of ovarian tissue or entire ovary for transplantation, followed by oocyte harvesting or natural fertilization, or collection of eggs and cryopreservation at different maturation stages is a new trend.

4.1 Pregnancy Rates and Calving

Recipient animals are subjected to ultrasonography 30 to 40 days after ET [155,156] or transrectal palpation at 30 to 60 days [9,15] after AI or ET to check the persistency of the corpus luteum present during the transfer of the embryos. Thereafter, confirmation of pregnancy is done by another transrectal palpation at least 90 to 180 days after the transfer.

Application of the advanced technologies particularly the AI, SOET/MOET, IVEP, OPU, cryopreservation, sperm sexing and cloning have resulted in the birth of several calves. The current success rate is 45 to 75% retrieval rate by ovum pick-up, 25 to 45% blastocyst development for in vitro embryo production, 25 to 55% percent pregnancies and 10 to 20% calving rate after embryo transfer. If genetically superior females will be used as oocyte donors before breeding them for natural gestation, their extra oocytes could be used to produce offsprings using surrogate mothers, thus, optimizing their reproductive performance and facilitating propagation of genetically superior animals.

5. Intracytoplasmic Sperm Injection (ICSI)

ICSI is a technology that involves collecting and injecting an acrosome and sperm-membrane-intact single sperm into the ooplasm of a metaphase II oocyte for fertilization [157]. ICSI has been used as an answer to male infertility in humans [158], whereas in livestock species the potential use of ICSI includes fertilization of oocytes with sex-selected sperms [159], circumventing the problems of polyspermy in cryopreserved oocytes (due to zona hardening and early extrusion of cortical granules) [160] and as an alternative procedure to pronuclear injection for producing transgenic animals using sperm as a vector to introduce genes [161]. Cryopreservation of oocytes results in zona hardening. This limits the penetration by sperm cells resulting in low success rates in conventional IVF. Similarly, motility of sex-sorted sperm cells is low and sperm concentration of sex-sorted sperm is limited Vis a Vis high cost of sorting and long sorting time. This makes the use of ICSI beneficial and practical.

To carry out the ICSI technique, the oocyte is held by vacuum against a micropipette with a bore diameter small enough to hold the oocyte and not suck it up. An individual sperm cell is then loaded into a microinjection pipette which is carefully guided into the oocyte using a micromanipulation machine, and is injected into the ooplasm of a metaphase II oocyte [162]. Many studies on ICSI in cattle have been performed [157] resulting in the live birth of calves [163]. It has been mentioned that artificial activation of bovine oocytes after conventional ICSI improved the fertilization rates and blastocyst development rates ranged from 6.1% to 40.1% [157]. Attempts to apply ICSI in the buffalo are recent [164-167]. The failure of male pronucleus formation into blastocysts generated after ICSI [168,169] has hampered the success of ICSI in buffalo. Diethyl Triol (DTT) treatment of sperm with reacted acrosomes before ICSI together with the activation of the ICSI oocytes overcame the problem and promoted successful male pronucleus formation in buffalo [168]. A recent report [167] informed of the successful production of blastocysts following ICSI in buffalo using immobilized live sperm and dead (killed by freezing) sperm. The motile spermatozoa were immobilized by scoring the sperm tail with the tip of the injection needle against the bottom of the dish immediately before injection while dead sperm was prepared by diluting the live sperm suspension 1:3 with TCM199+HEPES+ 10% FBS and directly freezing at -20°C. Dead sperm cells were thawed without any cryoprotectant at room temperature and used for ICSI [167].

6. Sperm Sexing

Sperm sexing provides the opportunity to produce offsprings of pre-determined sex. This was made possible by the creation and development of the flow cytometer. Proof of sorting efficacy has been demonstrated in many species and numerous applications in a variety of species is anticipated including endangered species and zoo and aquarium animals [170]. In water buffalo, successful results on sperm sexing were also achieved [13,14].

Accuracy of sexing is around 90% in most species. However, this technology is still rather expensive due to the low sexing efficiency where only about 20 million sperm cells are sorted in an hour which is the concentration required for 1 dose for artificial insemination in water buffalo. Furthermore, fertility of sexed sperm is lower than unsexed controls [84] leaving room for improvement.

Advances have been made in two main areas: increasing the number of sperm sexed accurately per unit time, and making the process less damaging to the sperm by optimizing the pressure in the flow cytometer [171].

Sperm sexing offers an obvious advantage in animal breeding programs. The possibility of choosing a male or female calf can be done by the use of X- and Y- sorted sperm in artificial insemination or in vitro fertilization programs. While studies and its application in water buffalo is limited, the techniques have been commercialized in the United States in the year 2005 with significant positive impact on dairy cattle and beef cattle production systems by producing the desired sex and nearly doubling the productivity [170]. In water buffaloes, calves of sex pre-determined were born [13]. The same authors found that buffaloes present about 3.8% differences in DNA contents between their X- and Y-chromosome-bearing spermatozoa, a difference large enough to allow successful sorting. Although the numbers of sorted spermatozoa per hour have currently attained larger figures than those from a decade ago (50–100 million compared to 350,000), these numbers imply fewer sperm doses for AI, reducing their application for conventional breeding. The technology is, however, very promising and provides opportunities for sex selection of IVEP-embryos [172,173], surpassing the need for sex diagnosis of the embryos (which is reliably done today by DNA probing, specific for the Y chromosome, but still time-consuming and – perhaps – not risk-free) [172,174]. Sex-sorting, albeit interesting for animal breeding strategies, is too costly (a flow sorter costs above $300,000 US), slow, and yields weak spermatozoa with a reduced lifespan [175,176]. Nevertheless, sexed semen is commercially available (male- or female- sorted spermatozoa) and it is becoming more competitive in cattle [177,178]. In buffaloes, deposition of sexed semen in the body of the uterus yielded higher pregnancy rates (45.5%) compared to when sexed semen was deposited in the uterine horn (32.3%) [179]. Conception rates did not differ between adult buffaloes and buffalo heifers; however there was a difference between bulls [179]. Most studies in buffaloes have reported the use of sexed semen for insemination after an estrus synchronization protocol [84,180-182]. A recent study, however, evaluated the insemination of 4521 swamp or crossbred buffaloes during natural estrus with X-sorted semen from river buffaloes and recorded a 48.5% pregnancy rate and 87.6% sex accuracy [183]. A similar previous study on Chinese swamp buffaloes (n=3863) that were inseminated with X-sorted river buffalo semen during natural estrus, recorded calving rates of 51.9% and sex accuracy of 89.0% [173]. These studies reflect the prospects of commercial application of insemination with sexed semen in the buffalo.

7. Cloning

7.1 Somatic Cell Nuclear Transfer

In SCNT, the nucleus of the somatic cell is transferred to an enucleated recipient cytoplast which is then stimulated to divide and develop into an embryo.

The potential benefits offered by this technology in the livestock industry are huge but the efficiency of implementation in water buffalo is slow and challenging. Successful cloning in water buffalo was shown through the production of cloned calves reported by the use of fetal fibroblasts or granulosa cells as donor cells [15], ear fibroblast nucleus from river buffalo fused into swamp buffalo oocyte cytoplasm [16], and in hand-made zona-free cloned vitrified embryos derived from enucleated oocytes reconstructed using adult skin fibroblast cells as nucleus donor [17,18].

Cloning can be done using a micromanipulator-based cloning procedure [184,185] or by hand-made procedure [186]. Studies on the use and efficiency of donor cell type [184,187-189] fusion, activation and culture systems [187,189] have been explored with varying success. Cloned embryos produced from nucleus transfer of buffalo fetal and adult somatic nuclei into enucleated bovine oocytes and subsequent development to the blastocyst stage has been reported [188,190]. Most previous studies have utilized adult or fetal fibroblasts as nucleus donors [184-187], however, some of the more recent reports depict the successful production of cloned embryos by nucleus transfer from cells other than fibroblasts such as somatic cells isolated from urine [191], amniotic fluid derived stem cells [192] and somatic cells from milk [193]. Cloned embryos have also been produced by nucleus transfer of somatic cells (fibroblasts from ear) from wild buffalo and oocytes from domestic buffaloes [194,195] and live births resulted following SCNT from river buffalo somatic cells and swamp buffalo oocytes [196]. Hand-made cloning has proved to be an efficient alternative to the conventional micromanipulator-based technique [186]. Research Vitro Cleaved medium was reported to be an effective culture system compared to mSOF and mCR2 medium for in vitro culture of zona-free cloned buffalo embryos reconstructed using adult skin fibroblast cells as nucleus donors [186]. Culturing zona-free cloned buffalo embryos on flat surfaces yielded significantly higher blastocyst rates than Well of the Wells (WOW) or microdrops (MD) [186].

The stage of the donor cell cycle is a major factor contributing to the success of SCNT in mammals [197]. To facilitate the remodeling and reprogramming of somatic cell nuclei, non activated metaphase II oocytes with high levels of maturation-promoting factor are generally employed as recipient cytoplasts for SCNT [198]. Donor cells must be arrested at the G1 or G0 stage of the cell cycle to avoid chromosomal damage and abnormal ploidy in the resulting embryos [15]. Serum starvation or use of reversible inhibitors of mammalian DNA polymerases that can block the cell cycle at the transition from the G1 to S phase are methods used to arrest cell lines in the G0 phase of the cell cycle [199].

7.2 Embryo Splitting

In embryo splitting, before reaching the differentiation stage an embryo is split in to 2, 3 or 4 depending on the efficiency of splitting. The separated blastomeres are further cultured for development to preimplantation stage. The resultant embryos are clones. The clone carries exactly the same DNA of the original embryo that was split. Embryo splitting is a technology that allows the development of several embryos from a single embryo. It is a cloning method where a healthy embryo is divided by taking some blastomeres and implanted into an empty zona pellucida to develop another embryo. It involves splitting or dissecting young embryos into several sections [200], usually at the 2 to 8 cell stage, although some reports used morula-stage embryos before the cells differentiated. The sections are further cultured to develop into a new embryo that upon reaching the preimplantation stage can be cryopreserved or transferred to recipient animals. The procedure requires specific microscopy and micromanipulations to carry out the extremely delicate micro-dissection needed when working with the embryo. A pregnancy rate of 120% was reported by this technique because by dissection, a 50 to 60% success rate after transfer of each half embryo was achieved when transfers were done using the same receptor female [200].

The earliest report of the first successful embryo splitting was achieved in the mouse [201].The researchers investigated the developmental potential of blastomeres from early preimplantation embryos. Other studies showed that 65% of half-embryos developed to term when split at the 2-cell stage and transferred to recipient females [202]. Colorado State University and Louisiana State University in the US developed this procedure. In water buffalo, Zhang et al., [19] succeeded in demonstrating the feasibility of using embryo splitting in combination with multiplex-nested PCR sexing to produce offsprings of controlled sex in swamp buffalo. They observed that grade-1 blastocysts yielded more viable demi-embryos than grade-2 and grade-3 blastocysts (P < 0.01; 73/92 (79.67%) vs 32/76 (47.05%) vs 26/94 (26.53%), respectively) where transplantation of the presumed-Y demi-embryo derived from a grade-1 blastocyst into a recipient resulted in the birth of a male buffalo calf.

Research initiatives on embryo splitting in water buffalo are limited. The invasiveness of the procedure and technical demands limit the use of this technique in buffaloes.

8. Embryo Sexing and Diagnosis

Embryo sexing and diagnosis allows one to know the sex of the product or to test for known genetic diseases of the embryo prior to its transplantation into a recipient. This can be done using techniques such as polymerase chain reaction (PCR), fluorescence in situ hybridisation (FISH) and karyotyping [203]. Preimplantation genetic diagnosis (PGD) allows the screening of embryos before transfer. The procedure begins with aspiration of two to three blastomeres from each embryo and a three hour DNA analysis to determine the sex and possible abnormalities. This technique uses DNA obtained from 1 or 2 blastomeres from demi embryos of 2 to 4 cells up to the blastocyst stage [204,205]. The sex determination assay involves using a primary (BRY.I), nested (BuRYN.I) and multiplex (BuRYN.I, ZFX/ZFY) PCR where the BRY.I and BuRYN.I primers are targeted to amplify Y-specific sequences, while the ZFX/ZFY loci were amplified to serve as a positive controls for both male and female samples [206]. The sex determination assay is accurate and is a highly reliable method for sexing buffalo embryos. More recently, a novel male-specific DNA sequence, cloned from Taiwan water buffalo genomic DNA templates, using random amplified polymorphic DNA (RAPD-PCR) was identified [207]. This resulted in the design of two primers, BuSexOPC16-F and –R that could amplify the male-specific fragment used in PCR for sexing. This technique was tested in Taiwan water buffalo genomic DNA samples and also in Taiwan yellow, Holstein, Angus, and Hereford cattle samples. It was demonstrated that the sex of these five breeds could be easily and effectively determined using the PCR technique. The loop-mediated isothermal amplification (LAMP) method is a novel method that amplifies a target DNA sequence specifically under isothermal conditions. LAMP has been developed for the rapid sexing of buffalo embryos through the identification of Y-chromosome-specific sequences [208]. When the target DNA is amplified by LAMP a white precipitate is formed [209] making the diagnosis easy. The total time required for sexing embryos using this approach was only 1 h [210] and the accuracy of sexing was 100% [211].

Embryo sexing is a potential spin-off of the embryo splitting technique given that a portion of the sectioned blastomeres can be sexed and genetically tested while the others can be cultured for further splitting and development of a sex-predetermined, pre-implantation embryo. This technique would guarantee the production of embryos of known sex and of their genetic quality.

9. Stem Cell Technology and Transgenesis

The production of embryos in vitro led to the discovery of stem cells technology in 1998 by Dr. James Thompson of University of Wisconsin-Madison. Stem cell technology offers a huge application on the potential treatment of human diseases such as cancer, Parkinson’s, Alzheimer’s, paralysis and even severe skin problems. Stem cells are undifferentiated cells that can proliferate and transform themselves to form a tissue which could be used to repair another tissue or provide new desired cells [212]. The embryo at morula stage and the inner cell mass of the blastocyst has undifferentiated blastomeres. These cells are stem cells and have the ability to form any of the terminally differentiated cells of the body. Studies on isolated embryonic stem cells (ES cells) from in vitro-produced buffalo embryos [213,214] demonstrated the capability of ES cell lines established from parthenogenetically produced buffalo embryos to differentiate in vitro and in vivo into a variety of cell types. The potential application of buffalo embryonic stem cells on human regenerative medicine is still to be revealed, however, with the advancement of molecular and nanotechnology, it may be possible that in vitro embryo production could find its way into the revolution of organ repair [215].

Development of stem cell technology in buffaloes finds application in the targeted genetic modification of this species. The testis has emerged as a source of pluripotent stem cells and some valuable information have been gathered [216,217]. Spermatogonia-specific proteins expressed in prepubertal buffalo testis and their utilization for isolation and in vitro cultivation of spermatogonia has been reported [218]. A few reports depict the production of other types of stem cells in buffaloes [219,220].

Production of transgenic animals is another breakthrough in reproductive biotechnology. It is the introduction of foreign genes into a livestock genome that allows the production of biomolecules and therapeutically relevant proteins for nutriceuticals or oral vaccines for humans using transgenic animals as bioreactors. This technology is called “biopharming”. The production of transgenic animals that produce milk with high similarity to human milk is a promising trend. Production of a transgenic buffalo calf is still a challenge. Production of cloned and transgenic embryos using buffalo embryonic stem cell-like cells isolated from in vitro fertilized and cloned blastocysts has resulted in the birth of a calf [105]. Similarly, the prospects of mammary gland specific expression vectors for high level of human insulin expression in transgenic buffaloes were evaluated [221] researching the possible use of molecules of buffalo origin in humans. However, the public’s perception of the science involved in transgenic animal production remains an issue that needs to be resolved besides the need for improvement on the technical aspect.

10. Molecular Genomics and Nanotechnology

Molecular genomics is the study of the structure, function and interactions of genes at the molecular level. It is the identification of the genes that control or express a certain trait and alteration of the animal genome, to create transgenic organisms that express a gene of interest. Apart from exploring complex traits, molecular genomics also offer enormous potential to improve the emerging reproductive biotechnologies in the buffalo by elucidating, unraveling and understanding the molecular basis of transcriptional control during the development of the preimplantation embryo which is of the utmost importance in the diagnosis and elimination or reduction of abnormalities associated with in vitro-produced embryos [222]. Its application may elucidate key molecular developmental dynamics in embryos resulting from somatic cell nuclear transfer to enable better understanding of the causes of the low efficiency of buffalo cloning.

There are new technological trends that emerged along with reproductive biotechnologies. Nanotechnology is believed to be a new trend that unveils gametes in minute details [157]; a sperm head colliding with the zona-pellucida shall be filmed, the radial blast of cortical reaction shall be known and all chemical messengers may be unveiled at nano-level. This technology may allow sending nano microprocessors into the animal’s womb to send signals that enable the control and prevention of diseases and other regulations that may improve the success and efficiency of the application and implementation of reproductive biotechnologies.