Assisted reproductive techniques in mares

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1 Received: 01 March 2018 Revised: 13 May 2018 Accepted: 16 May 2018 DOI: /rda PLENARY Assisted reproductive techniques in mares Katrin Hinrichs Department of Veterinary Physiology and Pharmacology, College of Veterinary Medicine & Biomedical Sciences, Texas A&M University, College Station, Texas Correspondence Katrin Hinrichs, Department of Veterinary Physiology and Pharmacology, College of Veterinary Medicine & Biomedical Sciences, Texas A&M University, College Station, TX Funding information American Quarter Horse Foundation; Clinical Equine ICSI Program, Texas A&M University; Link Equine Research Endowment Fund, Texas A&M University Abstract A wide variety of assisted reproductive techniques (ARTs) are available to aid in managing aspects of equine reproduction. Embryo recovery and transfer can be used to obtain more than one foal per mare per year, and to obtain foals from mares that cannot carry a foal to term. Oocyte recovery and either transfer to the oviduct of an inseminated recipient mare (oocyte transfer), or intracytoplasmic sperm injection (ICSI) and embryo culture can be used to obtain foals from mares with some types of subfertility, such as problems of the tubular tract. ICSI can be used to obtain foals when sperm number or quality is low. Because of its ease of use for the mare owner and efficiency, oocyte recovery and ICSI is being used in some cases for management of normally fertile mares and stallions. Oocytes can be recovered from live mares by the referring veterinarian, and shipped overnight to a laboratory for ICSI, without any decrease in oocyte or embryo viability. In case of unexpected death of a mare, ovaries or oocytes can be transported to the ICSI laboratory for production of embryos. Embryos produced both in vitro and in vivo can be biopsied to determine their genetic makeup before they are transferred. Equine embryos can be vitrified successfully; collapse of the blastocoele cavity allows efficient vitrification of expanded blastocysts. In contrast, cryopreservation of unfertilized equine oocytes still has low success. Genetics of valuable animals can be preserved via nuclear transfer (cloning) and several commercial companies offer this service clinically. KEYWORDS cloning, cryopreservation, embryos, horses, intracytoplasmic, organism, preimplantation diagnosis, sperm injections 1 EMBRYO TRANSFER Embryo transfer (ET), that is, recovery of an embryo from the uterus of a donor mare and transfer of the embryo to the uterus of a recipient mare, is a well- established equine assisted reproductive technique (ART) (Squires, McCue, & Vanderwall, 1999). The standard protocol for equine ET is to monitor the preovulatory follicle of the donor mare via ultrasonography per rectum. The mare is inseminated when the follicle appears to be ready to ovulate, or in coordination with induction of ovulation by administration of a GnRH or LH analog. The preovulatory follicle is monitored daily after insemination so that the day of ovulation (Day 0) can be determined. The embryo is flushed from the donor mare s uterus, typically on Day 7 or 8 after ovulation. Flushing is performed by filling the uterus with ~1 L of flush solution, then draining the fluid through a filter and repeating this multiple times. The embryo is located in the fluid retained in the filter, and is washed and transferred transcervically to the uterus of a recipient mare that ovulated from a day before to 3 days after the donor mare. With a fertile donor mare, the expected embryo recovery is about 75% and the expected pregnancy rate after transfer about 75%, for a ~50% recipient pregnancy rate per cycle (Squires et al., 1999; Vogelsang, Bondioli, & Massey, 1985), although experienced practitioners may report higher rates than this. Embryo transfer can be used to obtain more than one foal per donor mare per year, or to obtain foals from donor mares that the owner does not want to be pregnant (because the mare is in Blackwell Verlag GmbH wileyonlinelibrary.com/journal/rda Reprod Dom Anim. 2018;53(Suppl. 2):4 13.

2 HINRICHS 5 competition, the mare is too valuable to risk the potential dangers of pregnancy or foaling, or the mare has other issues, such as lameness, pelvic damage, or even maternal behavior). Embryo transfer can also be used to recover embryos from donor mares that are capable of conception but which suffer from repeated early embryonic death associated with factors such as uterine periglandular fibrosis or other age- related uterine changes (Kenney, 1978). The main limitations of embryo transfer as an equine ART are that (a) recovery of embryos requires that the donor mare have a functional tract, at least to the point that sperm can reach the oviduct, the mare ovulates normally, and the oviduct and uterus can support fertilization and development of the embryo until Day 7 or 8 after ovulation; (b) breeding the donor mare requires a full insemination dose of good- quality semen; and (c) superovulation of mares is problematic (Squires & McCue, 2007), so only the potential embryo resulting from ovulation of the one dominant follicle (or sometimes two, if the donor mare spontaneously double- ovulates) is available. Use of ET in mares in competition can be complicated by the effects of stress (Campbell, 2014), and repeated manipulation of the mare s uterus (breeding, uterine flush) can be associated with induction of endometritis (Campbell, 2014). 2 OOCYTE RECOVERY AND INTRACYTOPLASMIC SPERM INJECTION Oocyte recovery and intracytoplasmic sperm injection (ICSI) can be used for production of foals from mares that cannot become pregnant or provide an embryo under standard reproductive management. This includes mares with cervical, uterine or oviductal abnormalities or disease. However, ICSI will NOT be helpful if the cause of the mare s subfertility is due to her oocyte quality. In the clinical ICSI program at Texas A&M, the main driver for ICSI is on the part of the stallion owner, to allow breeding of mares by top stallions that have aged or died and so have only limited supplies of frozen semen available. The efficiency of frozen semen is greatly increased by ICSI, as only a few sperm are needed, from which one is selected to fertilize each oocyte. Frozen semen can be thawed, diluted up to 1:200, and refrozen in a large number of straws of ICSI doses (Choi, Love, Varner, & Hinrichs, 2006a) and used for ICSI successfully. If motility is good, a portion of this straw (a cut ) can be used at a time, for fertilization of numerous oocytes, thus allowing an existing store of frozen semen to produce a large number of embryos. 2.1 Oocyte recovery from live mares Mare management There are two main approaches to obtaining oocytes from the donor mare: aspiration of the one dominant, stimulated follicle to recover an in vivo-matured oocyte just before it ovulates; or aspiration of all the immature follicles on the ovaries, without ovarian stimulation, to recover immature oocytes that must then be matured in vitro. In oocyte terms, mature refers to an oocyte that has resumed meiosis and has reached metaphase II, the stage at which the oocyte is ovulated and at which it is capable of being fertilized. The oocyte resumes meiosis in response to either the endogenous LH signal, or after exogenous stimulation with a gonadotrophic hormone (GnRH analog or LH analog). For most practices, aspiration of immature follicles is the more efficient method for recovering oocytes for ICSI. A detailed description of materials and procedures for follicle aspiration has been presented previously (Hinrichs, 2010) so here I will discuss general considerations Aspiration of dominant, stimulated follicles There are several advantages to recovery of the mature oocyte from the dominant stimulated follicle (DSF), especially for the practitioner just starting to work with follicle aspiration. Compared to aspiration of immature follicles, aspiration of the DSF is easier because of: (a) the large follicular size; and (b) a high recovery rate (~80%; Carnevale & Ginther, 1995; Hinrichs, Matthews, Freeman, & Torello, 1998) as the oocyte cumulus is expanded due to gonadotropin stimulation, and thus the cumulus-oocyte complex is detached from the follicle wall. Oocytes recovered from the DSF have high developmental competence (ability to yield a viable embryo). One study reported a 70% blastocyst rate per injected DSF oocyte after ICSI, vs. ~30% blastocysts for in vitro- matured oocytes (Foss, Ortis, & Hinrichs, 2013); others have reported rates of 40% vs. 33% for the two types of oocytes (Jacobson, Choi, Hayden, & Hinrichs, 2010). However, recovery of the oocyte from the DSF has many disadvantages. It requires frequent monitoring of follicle growth and accurate timing of hcg/gnrh treatment to stimulate the dominant follicle. ICSI must be performed at a set time after gonadotropin stimulation (typically hr), as the gonadotropin stimulation starts meiotic resumption. Thus, the timing of gonadotropin stimulation must be coordinated with the ICSI laboratory. Because superovulation of mares is problematic, and presence of multiple large follicles on the ovaries make them difficult to manipulate (Maclellan et al., 2002), typically only the one preovulatory follicle (or sometimes two, if mare raises two DSFs) is available per aspiration procedure. Oocyte recovery from DSF may be higher if both hormones (hcg and GnRH) are administered for follicle stimulation, at least in old mares (Riera, Roldan, Gomez, & Hinrichs, 2016). Aspiration must be done when follicle has responded to the gonadotropin, to ensure expansion of the cumulus, but before ovulation; typically the DSF is aspirated hr after gonadotropin stimulation. In one study, recovery rates were highest when DSF were aspirated hr after hcg + deslorelin treatment (Riera et al., 2016). If oocytes will be shipped to the ICSI laboratory, recovering the oocyte at this time allows ~12 hr to transport the oocyte to the laboratory.

3 6 HINRICHS Aspiration of immature follicles If oocytes are to be recovered from immature follicles, aspiration can be done on a predictable set schedule, e.g., once every 14 days, without having to follow ovarian activity between aspirations (Jacobson et al., 2010). Alternatively, donor mares can be monitored by ultrasound to select a time for follicle aspiration when an adequate number of small follicles (e.g., 8 10), and no large preovulatory follicle, are present (Galli, Duchi, Colleoni, Lagutina, & Lazzari, 2014). Typically, all follicles on the ovary over ~5 8 mm diameter are aspirated; the number of oocytes recovered depends upon the number of follicles present. The population of mares that come to our facility are typically old or subfertile, and many have only a few follicles developing at any time. Different breeds of mares may tend to have different follicle numbers, and may have different rates of embryo production after ICSI (Galli et al., 2014). Aspiration of immature follicles can be performed year- round; although follicles are smaller in the non- breeding season, the number of follicles present does not differ from that in the breeding season (Choi et al., 2016; Hinrichs & Schmidt, 2000) and normal maturation and blastocyst rates can be obtained from oocytes recovered in the non- breeding season (Choi et al., 2016). The oocytes recovered from the immature follicles are in prophase of meiosis, a resting phase they can be handled at room temperature and can be held overnight at room temperature in simple holding medium with no effect on viability or resulting blastocyst development (Choi, Love, Varner, & Hinrichs, 2006b; Foss et al., 2013). Because of this, there is a large leeway in the timing of working with immature oocytes they can be placed in maturation right away or can be held overnight or even potentially up to two nights (Foss et al., 2013) before being put into maturation, thus allowing ICSI to be scheduled at a convenient time. The main disadvantages to aspiration of immature follicles are that the aspiration procedure is more difficult the follicles are small and more difficult to hit with the needle, and because the cumulus has not yet expanded, the cumulus- oocyte mass is tightly attached to the follicle wall (Hawley, Enders, & Hinrichs, 1995). This makes it necessary to repeatedly fill and empty the follicle and scrape the follicle wall with the needle to obtain a reasonable recovery rate (~50%). The oocyte that is recovered is immature, and must undergo in vitro maturation (IVM). Because some of these oocytes were recovered from juvenile follicles and some from atretic (dying) follicles, only about 60% of the recovered oocytes will mature in culture, and thus be fertilizable (Hinrichs et al., 2005), and the ability of the oocytes to produce embryos after ICSI is lower than that for the in vivo- matured oocyte from the DSF. Despite the above disadvantages of aspiration of immature follicles, the increase in number of oocytes recovered when these follicles are used results in a much higher rate of blastocyst development per aspiration than does aspiration of the DSF alone (~1 blastocyst per aspiration vs blastocysts per aspiration, respectively; Jacobson et al., 2010). It is possible to aspirate immature follicles at the same time as a DSF follicle. The fact that the immature follicles experienced hcg and/or GnRH hr previously does not appear to affect the maturation or blastocyst rate with these oocytes (unpublished observations). However, two ICSI procedures will have to be performed in this case, as the DSF oocyte will be mature and ready for ICSI a day earlier than are the oocytes recovered from immature follicles and then matured in vitro Follicle aspiration technique It is possible to aspirate the DSF by placing a needle through the flank, a method that is simple and requires only the most basic materials (Hinrichs et al., 1998). However, most practitioners prefer to be able to visualize the follicle and so use transvaginal ultrasoundguided follicle aspiration (TVA). With this technique, the ultrasound probe is placed in a long holder that has a channel within it to guide the needle. The ovary is grasped per rectum and manipulated to bring the ovary to the peritoneal side of the vaginal wall. A needle (typically a 12- ga double- lumen needle) is passed through the vaginal wall into the follicle and the contents of the follicle aspirated. As noted above, if the follicle is immature, it is necessary to collapse the follicle and scrape the needle against the follicle wall repeatedly to dislodge granulosa cells, and hopefully the cumulus- oocyte complex. We do not administer antibiotics routinely to our research mares (which undergo TVA essentially every 14 days year- round) to prevent establishment of antibiotic- resistant bacteria in the herd. We evaluated the effect of repeated TVA on mare health and ovarian morphology in our research mares, and found that the incidence of complications was rare (1 case in more than 300 TVAs; Velez et al., 2012). However, complications such as ovarian abscess, peritonitis and death are possible after TVA (Bøgh, Brink, Jensen, Lehn- Jensen, & Greve, 2003; Vanderwall & Woods, 2002; Velez et al., 2012). For these reasons, and because client mares typically have a limited number of TVAs per year and do not all live in the same environment, we administer a broad- spectrum antibiotic to client mares after TVA. It is important that mare owners are aware that transvaginal follicle aspiration carries risks for the donor mare; it is not a benign procedure. The transvaginal probe is exposed to bacteria as it passes through the vestibule (which has heavy bacterial contamination; Hinrichs, Cummings, Sertich, & Kenney, 1988) when it is inserted into the vagina. Thus, the needle transits through a potential non- sterile environment repeatedly as it is extended from the probe handle through the vaginal wall into the peritoneum and ovary. Attempts at sterilizing the vagina, e.g., with dilute betadine, should not be attempted as antimicrobials are toxic to oocytes. 2.2 Handling and packaging oocytes Few practices have the capability to perform equine ICSI efficiently; thus typically oocytes are shipped to central ICSI facilities. Handling and shipping of maturing oocytes from aspiration of DSF is much different from that for immature oocytes. Since oocytes from DSF are in the process of meiosis, they need to be kept at body temperature until the time of ICSI, and so should be shipped in a portable

4 HINRICHS 7 incubator. The oocyte is actively metabolizing, so it is assumed that it will need a culture medium (e.g., M199 with 10% fetal bovine serum) rather than a simpler embryo holding medium, but no data is available on this. An adequate volume of medium must be provided, to avoid the medium becoming acidic due to the metabolism of the oocyte and its large cumulus. We put these cumulus- oocytes complexes into about 5 ml of medium for shipment. Immature oocytes may be handled at room temperature, and then held overnight, or shipped overnight, in media at room temperature (Choi et al., 2006b; Diaw, Salgado, Canesin, Gridley, & Hinrichs, 2018). A variety of commercial embryo holding media have been used for this with success (Diaw et al., 2018; Foss et al., 2013). The temperature of holding/shipment should be room temperature (~22 C). If the overnight holding temperature is too warm (30 C), oocytes will start to mature in the holding medium and lose viability (Martino et al., 2014); if the oocytes are held at cold temperatures (4 7 C) the maturation rate and blastocyst rate are significantly lowered (Diaw et al., 2018). A passive cooling device such as an Equitainer (Hamilton Biovet LLC, Ipswich, MA, USA) with all materials at room temperature, or a container designed to hold at room temperature (EquOcyte, Hamilton Biovet) work well for this purpose. 2.3 Postmortem ovary and oocyte recovery and shipment for ICSI When a mare dies unexpectedly, the ovaries can be removed from the mare and shipped to an ICSI laboratory with resulting production of embryos and foals (Hinrichs et al., 2012). If the ovaries are to be shipped, they should be allowed to cool slowly to room temperature during shipment, and potentially to temperatures of ~12 C (Preis et al., 2004), but no lower. Equitainers or EquOcyte containers work well for ovary shipment. The ovaries should be shipped by the fastest means possible to the laboratory; best results are thought to be obtained if the ovaries are received within 6 hr of the mare s death. Alternatively, the ovaries can be processed at the mare s location or at a laboratory within a few hour s transport, to recover immature oocytes, and the oocytes shipped to the ICSI laboratory as for immature oocytes recovered from live mares. This relieves the time pressure on getting the material to the ICSI laboratory. Recovery of oocytes from follicles in the isolated ovary requires scraping of the granulosa cell layer to dislodge the oocyte from the follicle wall, as is done during TVA of immature follicles. If the recovery rate is not extremely important, this can be done by scraping the follicle with a needle attached to a vacuum, either within the follicle as it is rinsed with medium from another syringe (Smits et al., 2009), or by opening the follicle with a scalpel and scraping the surface with the needle and rinsing the needle with medium (Canesin et al., 2017). However, in the case of a valuable mare that has died, recovering every possible oocyte from the ovaries is essential. A high recovery (mean, 18 oocytes/mare; range, 0 35; Hinrichs et al., 2012) can be achieved by opening follicles with a scalpel and scraping the wall with a bone curette (Hinrichs & DiGiorgio, 1991). Embryo holding medium may be used to wash the collected cells into a Petri dish. The wall is then scraped again and those cells washed into the dish. This is repeated until the entire surface of the follicle has been scraped. We use one Petri dish (top or bottom) per follicle. The oocytes are located in the collected cells and then handled as for immature oocytes collected by TVA. In a series of 16 client mares in which oocyte recovery and ICSI was done post- mortem, 10 foals were produced (Hinrichs et al., 2012). 2.4 The ICSI laboratory While conventional in vitro fertilization is not yet repeatable in the horse, intracytoplasmic sperm injection (ICSI) can be used for in vitro equine embryo production. An effective method for in vitro culture of equine ICSI- derived zygotes to the blastocyst stage was reported in 2005 (Hinrichs et al., 2005), and the first report of clinical application of equine ICSI was in 2007 (Colleoni et al., 2007). Since that time, equine ICSI has become a widespread procedure. At the ICSI laboratory, oocytes are incubated to either complete maturation (oocytes from DSF) or to initiate and complete maturation (oocytes from immature follicles). Maturation in our laboratory is performed in a cell culture medium (M199) with 10% fetal bovine serum and 5 mu/ml FSH, under 5% CO 2 in air; other laboratories may use different media. Galli, Colleoni, Duchi, Lagutina, and Lazzari (2007) found a higher blastocyst rate when a DMEM/F- 12 based maturation medium was used; however, we have found the opposite, i.e., a marked reduction in both maturation and blastocyst production with DMEM/F- 12 based maturation medium (unpublished results). These conflicting results may reflect differences in the specific makeup (additives, ph, osmolarity, etc.) of the media and in the culture environments used. Oocytes are typically matured for hr (Hinrichs et al., 2005) then the cumulus is denuded and those oocytes showing a polar body are selected for ICSI. Sperm for ICSI are prepared by swim up, direct wash, or density- gradient or single- colloid centrifugation; little information is available on the effect of sperm preparation method on equine ICSI outcome. ICSI is performed via micromanipulation on an inverted microscope. Sperm motility is slowed by placement in solution of a polyvinylpyrrolidone (typically 7% 10%) and if possible, motile sperm with normal morphology are selected. The plasma membrane of the sperm is disrupted by manipulation with the injection pipette, and the sperm is injected into the cytoplasm of the oocyte. Equine ICSI may be performed with either a conventional pipette or with a Piezo- driven pipette; in a recent study we found no difference in blastocyst rates per injected oocyte between the two techniques (39% and 40%, respectively; Salgado RM, Brom-de-Luna JG, Resende HL, Canesin HS, Hinrichs K.). However, the quality of the embryos, as evidenced by nucleus number and nuclear fragmentation rate, was significantly higher in blastocysts produced via Piezo- driven ICSI. It is important that the practitioner work with an ICSI laboratory that has an established record of successful embryo and foal production. An effective ICSI laboratory should be able to provide you data on the number of clinical cases (oocyte collections) on which

5 8 HINRICHS ICSI was performed the previous year, the in vitro maturation rate achieved (prefer 50%), the blastocyst rate per injected oocyte (prefer 20%), the pregnancy rate after transfer of blastocysts to recipient mares (prefer 65%) and the foaling rate per established pregnancy (prefer 80%). The ICSI laboratory may have a herd of recipient mares, or may ship embryos back to the practitioner for transfer. We consider in vitro- produced blastocysts, regardless of the day they are observed in culture and transferred, to be the equivalent of early (Day- 6) in vivo- recovered blastocysts. Therefore, in vitro- produced blastocysts should be transferred to mares that are 4 6 days after ovulation on the day of transfer. Little information is available on the health of foals produced by ICSI. In a recent study (n = 10 per group), we found no significant differences in foal general health, size or weight, or in placental morphology or selected gene expression, among foals produced by natural conception, embryo transfer or ICSI (Valenzuela et al., 2017). 3 TRANSFER OF OOCYTES TO THE OVIDUCT OF INSEMINATED MARES (OOCYTE TRANSFER) In some locations, effective ICSI laboratories are not available. An alternative method for production of foals from oocytes recovered from donor mares is to perform oocyte transfer, that is, the transfer of mature oocytes to the oviducts of inseminated recipient mares. In this case, the oocyte is fertilized in the oviduct of the recipient mare, and the resulting embryo is carried to term by that recipient. Clinical oocyte transfer has been conducted successfully on a commercial basis (Carnevale, Coutinho da Silva, Panzani, Stokes, & Squires, 2005; Riera et al., 2016). The main drawbacks to oocyte transfer as a method for production of foals from isolated oocytes are (a) the transfer of the oocyte requires surgery on the recipient mare to expose the oviduct; (b) since the recipient mare is inseminated, a full insemination dose of good- quality semen is required; and (c) if multiple oocytes are available, then the decision must be made to either perform multiple surgeries or to transfer multiple oocytes to the oviducts of a single recipient. The latter option requires either reduction of any embryonic vesicles >1 as visualized on ultrasonography at ~14 days (Carnevale, Coutinho da Silva, Preis, Stokes, & Squires, 2004), or subsequent flushing of the uterus of the recipient mare to recover embryos, with associated potential for losses during recovery and transfer to a secondary recipient (Riera et al., 2016). For effective oocyte transfer, a mature (metaphase II) oocyte is necessary. This can be obtained after in vitro maturation of oocytes recovered from immature follicles; however, this results in multiple oocytes of variable developmental potential, thus presenting the multiple- oocyte quandary mentioned above. Nevertheless, in case of post- mortem oocyte recovery, in vitro maturation is the only possible approach, and these oocytes may be used successfully for oocyte transfer (Carnevale, Coutinho da Silva, Preis et al., 2004). Since the cumulus is thought to be important in allowing the oviduct to capture and retain the oocyte, the oocytes are transferred with the cumulus intact, thus it is not possible to know how many of the transferred oocytes are mature and thus fertilizable. When the goal is to produce a foal from a live donor mare, the most effective approach for use of oocyte transfer is to recover the in vivo- matured oocyte from the donor mare s DSF. This provides one mature oocyte, of maximum developmental competence, for transfer. The donor mare is managed as for recovery of the oocyte from the DSF, above. The recipient mare should also be in estrus or under estrogen influence, as her tract must be capable of correctly handling the sperm after insemination, and supporting fertilization of the transferred oocyte. After the transfer, the recipient mare must form a functional corpus luteum or be under progesterone influence, so that her tract allows normal embryo development and pregnancy. This can be assured either by choosing a recipient mare in estrus with a large preovulatory follicle (the recipient follicle is aspirated to remove the oocyte, to prevent conception from this oocyte (Carnevale & Ginther, 1995)) or using an anestrus or early estrus recipient that is treated with estrogen before the transfer and progesterone afterward (Hinrichs, Provost, & Torello, 2000). The transfer is performed via standing flank laparotomy (Hinrichs et al., 1998; Riera et al., 2016); after the peritoneum is opened, the ovary is grasped and brought to the incision. The infundibulum of the oviduct is attached to the area of the ovulation fossa in the mare, and so is easily identified. The oocyte is loaded into a narrow- gauge pipette, and the pipette placed into the ampulla of the oviduct, and the oocyte expelled. The ovary is replaced in the abdomen and the incision closed. The recipient mare is inseminated, typically before the oocyte transfer (Carnevale, Coutinho da Silva, Maclellan, Seidel, & Squires, 2004; Riera et al., 2016). Perhaps due to all the manipulations performed and drugs administered, the recipient mare appears to have an increased tendency to undergo post- breeding endometritis (Hinrichs, Betschart, McCue, & Squires, 2000) and should be monitored for this and treated if it occurs. When oocyte transfer is performed using oocytes from fertile mares, the pregnancy rate is typically >70% (Carnevale & Ginther, 1995; Carnevale, Coutinho da Silva, Maclellan et al., 2004; Hinrichs et al., 1998). However, the pregnancy rates are notably lower when the procedure is used clinically, probably related to the age and reproductive status of the donor mares (Carnevale et al., 2005; Riera et al., 2016). 4 EMBRYO BIOPSY FOR PREIMPLANTATION GENETIC DIAGNOSIS In vitro- produced (IVP) embryos, as well as embryos recovered from live mares, can be biopsied to allow preimplantation genetic diagnosis. Biopsy is performed via micromanipulation, by removing a small number of trophoblast cells with the micropipette. Embryos biopsied with a micropipette have normal embryo viability (Choi, Gustafson Seabury et al., 2010; Guignot et al., 2015; Herrera et al.,

6 HINRICHS ). Biopsy with a microblade is possible in smaller embryos (<300 μm diameter) but when used in larger embryos drastically lowered pregnancy rates (Guignot et al., 2015; Seidel, Cullingford, Stokes, Carnevale, & McCue, 2010; Troedsson et al., 2010). The cells recovered on biopsy can provide an accurate genetic diagnosis after whole genome amplification and PCR (Choi, Penedo, Daftari, Velez, & Hinrichs, 2015); equally high accuracy was found for biopsies from IVP and in vivo- derived embryos. Preimplantation genetic diagnosis allows embryos to be selected for sex, or allows embryos carrying disease- related mutations to be selected against. In vivo- recovered embryos can be shipped to the ART laboratory overnight, biopsied, and shipped back for immediate transfer with a resulting normal pregnancy rate (Choi, Gustafson Seabury et al., 2010). If the embryo is an expanded blastocyst, the biopsy procedure is more difficult due to the tough equine embryonic capsule, and the tighter adherence of the trophoblast cells to each other. Biopsy of expanded blastocysts causes loss of blastocoele fluid and collapse of the blastocyst, but these embryos recover within a few hours in culture, and provide a normal pregnancy rate (Choi, Gustafson Seabury et al., 2010). The main drawbacks are the need for micromanipulation, and the requirement for a functional genetics laboratory to perform the genetic determination. Ways to simplify the micromanipulation involved have been proposed, including aspiration of only blastocoelic fluid (Herrera et al., 2015), but have not yet been shown to be repeatable. Methods for simplified PCR techniques, that can be performed in practice, are currently under development but have not yet been validated. 5 EMBRYO CRYOPRESERVATION Equine embryos less than 300 μm in diameter can be frozen or vitrified successfully using techniques developed in other species. This includes IVP embryos (Colleoni et al., 2007) and small in vivorecovered equine embryos (Eldridge- Panuska, Caracciolo di Brienza, Seidel, Squires, & Carnevale, 2005; Hochi, Maruyama, & Oguri, 1996; Slade, Takeda, Squires, Elsden, & Seidel, 1985). However, obtaining small embryos in vivo necessitates recovery of embryos on Day 6 after ovulation. While the equine embryo has descended from the oviduct to the uterus by late Day 5 after ovulation (Freeman, Weber, Geary, & Woods, 1991), uterine flush on Day 6 has been reported to result in lower embryo recovery compared to that for Day 7 or 8 (Battut, Colchen, Fieni, Tainturier, & Bruyas, 1998; Iuliano, Squires, & Cook, 1985). Cryopreservation of expanded equine blastocysts (>300 μm) was initially problematic, associated with low pregnancy rates after either vitrification or freezing (Barfield, McCue, Squires, & Seidel, 2009; Eldridge- Panuska et al., 2005; Slade et al., 1985). When we evaluated methods for embryo biopsy (see above), it became evident that large expanded equine blastocysts can recover from blastocoele collapse. This led us to investigate whether blastocysts collapsed via micromanipulation could be successfully vitrified. We found that large expanded blastocysts, up to 600 μm in diameter, could be successfully vitrified after collapse. Using an ethylene glycol/sugar vitrification system, these embryos provided normal ongoing pregnancy rates after transfer (71%; Choi, Hartman et al., 2010). The media used affected results; embryos vitrified using an alternative medium containing dimethylsulfoxide provided similar initial pregnancy rates, but only 25% of established pregnancies went on to the heartbeat stage. Blastocoele collapse has been used with a different cryoprotectant system, incorporating media with ethylene glycol and glycerol and using the Cryolock open vitrification device (Diaz, Bondiolli, Paccamonti, & Gentry, 2016). This system provided good pregnancy rates (5/6) for Day- 8 in vivo- recovered embryos. The vitrification process itself is straightforward and rapid; the device carrying the embryo is directly plunged in liquid nitrogen. Further research is needed to develop a simple device that can safely collapse the blastocoele without need for a micromanipulator, allowing vitrification of expanded equine blastocysts in standard veterinary practice. 6 OOCYTE CRYOPRESERVATION Cryopreservation of equine unfertilized oocytes is still not efficient enough to recommend for clinical use. Two reports are available on embryo production from in vivo- matured oocytes that were vitrified and warmed (Maclellan, Stokes, Preis, McCue, & Carnevale, 2010; Maclellan et al., 2002). Vitrification of immature oocytes, followed by warming, maturation and ICSI, has resulted in moderate rates of maturation and embryo production (1% 15% blastocysts per injected oocyte; Tharasanit et al., 2006; Canesin et al., 2017; Ortiz- Escribano et al., 2017; Canesin et al., 2018). There is extensive interest in oocyte cryopreservation as a means to preserve valuable female genetics, store oocytes after unexpected death of a mare, or provide a supply of oocytes for research. Vitrification of in vivo- matured oocytes is successful in humans, yielding fertilization, blastocyst formation, pregnancy, and live birth rates equivalent to those for fresh oocytes (Cobo, Meseguer, Remohi, & Pellicer, 2010; Kuwayama, Vajta, Kato, & Leibo, 2005). Oocyte vitrification in domestic animals, especially of immature oocytes, is generally less efficient, possibly due to the larger amount of lipid present in these oocytes. Much further work is needed in this area. 7 CLONING (NUCLEAR TRANSFER) The first cloned horse foal was produced in Italy by Galli et al. (2003). Since then, production of cloned horse foals has been reported in the scientific literature by our group at Texas A&M (Hinrichs, Choi, Varner, & Hartman, 2007), two laboratories in Argentina (Gambini, Jarazo, Olivera, & Salamone, 2012; Olivera et al., 2016), and a laboratory in Korea (Lee et al., 2015). In total, around 75 cloned foals have been reported in the scientific literature. However, production of hundreds of cloned foals has been announced in the popular press by commercial companies, largely in the US and Argentina. The

7 10 HINRICHS production of cloned Polo horses in Argentina has been spurred on by the participation of well- known athletes and owners in this field; for example, renowned player Adolfo Cambiaso won the Palermo Open polo match in 2016 riding six clones of one of his best mares. While some cloned horses are being used in performance, the power of cloning is its use in breeding stock to preserve genetics of rare, aged or deceased horses, or allow production of foals from champions that were castrated before reproducing. The procedures used to perform nuclear transfer reflect the steps involved in fertilization. In both normal fertilization and in cloning, the metaphase II oocyte is essential, as it provides all of the cytoplasm that will form the early embryo, contains factors needed for processing of the sperm or somatic cell chromatin after entry and for oocyte activation, maternal mrnas for production of stagespecific proteins needed in the first cleavage stages, and all other components necessary for embryo development the oocyte can be considered to be an embryo in waiting. In normal fertilization, the role of the sperm is twofold: to contribute half the chromosomes for the embryo, and to stimulate the oocyte to progress from metaphase II and initiate embryo development ( activation ). Entry of the sperm into the oocyte triggers activation through release of the sperm factor, PLCζ. The PLCζ initiates a cascade that results in calcium release from the oocyte s internal stores (Saunders et al., 2002). The released calcium is then taken up again into the stores, released again, etc., resulting in oscillations of calcium in the cytoplasm, which causes breakdown of the cellular mechanisms keeping the oocyte in MII. The oocyte then completes meiosis, and starts to process the male and female chromatin to form pronuclei. The sperm chromatin is completely remodeled by the oocyte, which replaces the sperm protamines with histones, and other sperm chromatin proteins with maternal proteins, removes much of the sperm- specific DNA methylation, and repairs any DNA damage (review, Rivera & Ross, 2013). Within the pronuclei, the chromatin is duplicated, then the two pronuclei fuse, the nuclear membranes break down, the chromatin condenses on the first mitotic metaphase plate, and the first embryonic cell division occurs. In cloning, the two roles of the sperm (bringing in DNA and activating the oocyte) are replaced by the steps of nuclear transfer. In this case, the nuclear DNA of the oocyte is removed, and all of the chromosomes are brought in from outside, in the nucleus of the donor cell. The oocyte is then exogenously activated to trigger embryonic development, utilizing the same mechanism as for normal fertilization, i.e., by causing an increase in cytoplasmic calcium. This is done either by inducing calcium to enter the oocyte from the medium (e.g., by electrical pulse, ethanol treatment, or calcium ionophore) or, in our laboratory, by injecting sperm cytoplasmic extract, which contains sperm factors and induces calcium oscillations similar to those obtained after fertilization (Bedford, Kurokawa, Hinrichs, & Fissore, 2003). As in natural fertilization, the increase in intracytoplasmic calcium causes oocyte activation. The oocyte then processes the introduced chromatin, replacing somatic cell proteins with maternallyderived proteins and removing cell- specific DNA methylation (Wen, Banaszynski, Rosenwaks, Allis, & Rafii, 2014). A pronucleus (termed a pseudo- pronucleus in cloning) forms (Wakayama & Yanagimachi, 2001). The chromatin is duplicated within the pseudo- pronucleus, then the pronuclear membrane breaks down, the chromatin condenses on the first mitotic metaphase plate and the first embryonic cell division occurs. The efficiency of cloning and the health of the cloned foal depend greatly on the techniques used in the cloning laboratory. There are numerous methods for maturing the oocyte, preparing the donor cells (which must be at a stage of the cell cycle synchronized with the stage of the oocyte), enucleating the oocyte, combining the somatic cell with the enucleated oocyte, activating the recombined oocyte, and treating the recombined oocyte to enhance its ability to process the transferred DNA (Choi et al., 2013; Galli et al., 2003; Gambini, De Stefano, Bevacqua, Karlanian, & Salamone, 2014; Hinrichs et al., 2007; Lagutina et al., 2005; Olivera et al., 2016), all of which may affect how well the oocyte can do its job. In cloning, the job of the oocyte is to reprogram the transferred somatic cell chromatin so that the DNA functions as needed by an early embryo. In other species, notably sheep and cattle, cloned pregnancies tend to be lost throughout gestation, and cloned offspring may suffer from problems including Large Offspring Syndrome, organ abnormalities, or endocrine disease. Placental abnormalities are common. Little data is available on the overall health of foals produced by cloning. We reported findings on health of 14 cloned foals; two of the foals died in the 2 weeks post- partum, and 6 of the remaining foals (50%) exhibited some problem, including neonatal maladjustment syndrome, enlarged umbilical cord, and/or front leg contracture, all of which resolved with treatment (Johnson, Clark- Price, Choi, Hartman, & Hinrichs, 2010). Similar problems in cloned foals have been reported by others (Gambini et al., 2014; Olivera et al., 2016), the majority of which also resolved with treatment. These neonatal problems are also seen in normally- conceived foals, but cloned foals tend to have a higher incidence. Abnormalities of the placenta including placental edema, detachment from the uterus, presence of cystic structures, and placentitis have also been reported in equine cloned pregnancies (Lagutina et al., 2005; Pozor et al., 2016). Recently, work from Argentina has suggested that use of bonemarrow derived mesenchymal stem cells as donor cells for nuclear transfer may increase the likelihood of production of normally healthy foals (Olivera et al., 2016, 2018). The possibility exists that cloning could be performed using somatic cells isolated from frozenthawed semen, which would allow production of clones in cases in which frozen semen was the only existing source of cells from an individual. One live calf has been produced in this manner (Selokar et al., 2016). Somatic cells have been successfully isolated from frozen- thawed equine semen (Brom- de- Luna, Canesin, Wright, & Hinrichs, 2018) but cloning was not attempted. Cloned foals have been produced from oocytes recovered from immature follicles by TVA (Choi, Velez, Macias- Garcia, & Hinrichs, 2015; Choi et al., 2013) including production of a mitochondrial- identical cloned foal via recovery of oocytes from two mares maternally related to the donor horse. Currently, few breed registries allow registration of cloned horses, however, many competitions that do not require strict breed

8 HINRICHS 11 registry allow cloned horses to compete, including the Fédération Equestre Internationale (FEI), which oversees equestrian competitions worldwide, including the Olympics. ACKNOWLEDGEMENTS Work in the author s laboratory was supported by the American Quarter Horse Foundation, the Clinical Equine ICSI Program, Texas A&M University, and the Link Equine Research Endowment Fund, Texas A&M University. CONFLICTS OF INTEREST The author has no conflicts of interest to declare. REFERENCES Barfield, J. P., McCue, P. M., Squires, E. L., & Seidel, G. E. Jr (2009). Effect of dehydration prior to cryopreservation of large equine embryos. Cryobiology, 59, cryobiol Battut, I., Colchen, S., Fieni, F., Tainturier, D., & Bruyas, J. F. (1998). Success rates when attempting to nonsurgically collect equine embryos at 144, 156 or 168 hours after ovulation. Equine Veterinary Journal Supplement, 25, Bedford, S. J., Kurokawa, M., Hinrichs, K., & Fissore, R. A. (2003). 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