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BOR - Papers in Press, published online ahead of print August 6, 2003.
Biol Reprod 2003, 10.1095/biolreprod.103.018515
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BIOLOGY OF REPRODUCTION 69, 1895–1906 (2003)
DOI: 10.1095/biolreprod.103.018515
© 2003 by the Society for the Study of Reproduction, Inc.

Embryo

Effects of In Vitro Production on Horse Embryo Morphology, Cytoskeletal Characteristics, and Blastocyst Capsule Formation

Jordi L. Tremoleda1,3, Tom A.E. Stout3, Irina Lagutina5, Giovanna Lazzari5, Mart M. Bevers2,4, Ben Colenbrander3,4, and Cesare Galli5

Departments of Equine Sciences3 Farm Animal Health,4 Faculty of Veterinary Medicine, Utrecht University, 3584 CM Utrecht, The Netherlands Laboratorio di Tecnologie della Riproduzione,5 I.S.I.L.S., Cremona 26100, Italy


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Blastocyst formation rates during horse embryo in vitro production (IVP) are disappointing, and embryos that blastulate in culture fail to produce the characteristic and vital glycoprotein capsule. The aim of this study was to evaluate the impact of IVP on horse embryo development and capsule formation. IVP embryos were produced by intracytoplasmic sperm injection of in vitro matured oocytes and either culture in synthetic oviduct fluid (SOF) or temporary transfer to the oviduct of a ewe. Control embryos were flushed from the uterus of mares 6–9 days after ovulation. Embryo morphology was evaluated with light microscopy, and multiphoton scanning confocal microscopy was used to examine the distribution of microfilaments (AlexaFluor-Phalloidin stained) and the rate of apoptosis (cells with fragmented or terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling-positive nuclei). To examine the influence of culture on capsule formation, conceptuses were stained with a monoclonal antibody specific for capsular glycoproteins (OC-1). The blastocyst rate was higher for zygotes transferred to a sheep's oviduct (16%) than for those cultured in SOF (6.3%). Day 7 IVP embryos were small and compact with relatively few cells, little or no blastocoele, and an indistinct inner cell mass. IVP embryos had high percentages of apoptotic cells (10% versus 0.3% for in vivo embryos) and irregularly distributed microfilaments. Although they secreted capsular glycoproteins, the latter did not form a normal capsule but instead permeated into the zona pellucida or remained in patches on the trophectodermal surface. These results demonstrate that the initial layer of capsule is composed of OC-1-reactive glycoproteins and that embryo development ex vivo is retarded and aberrant, with capsule formation failing as a result of failed glycoprotein aggregation.

assisted reproductive technology, early development, gamete biology


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
A variety of assisted reproductive techniques, including conventional in vitro fertilization (IVF), intracytoplasmic sperm injection (ICSI), oocyte transfer, and nuclear transfer, have been performed or attempted with equine gametes [1–5]. However, the relatively low efficiency of the in vitro culture (IVC) stages has limited their commercial application in this species. In particular, the current techniques for in vitro oocyte maturation, conventional IVF, and culture of zygotes to the blastocyst stage are inadequate. For example, although only two foals have been produced via conventional IVF [1, 6], the adoption of ICSI [3, 7] has, by circumventing the need for sperm to penetrate the zona pellucida (ZP), resulted in the birth of several foals [3, 8–11]. Still, most ICSI pregnancies have resulted from transferring immediate postcleavage zygotes to the oviduct of recipient mares [3, 8, 9], whereas only a few have been produced via IVC to the blastocyst stage and transfer to the uterus [4, 10, 11]. In support of the apparent adverse effect of culture, McKinnon et al. [9] reported that even incubation for 26–28 h between ICSI and oviductal transfer resulted in a lower pregnancy rate than transfer of presumptive zygotes 4–8 h after ICSI. Thus, if a viable, complete horse embryo in vitro production (IVP) system is desired, and on grounds of cost and recipient welfare it is, one critical step is to devise a culture system that supports embryo development to a stage suitable for uterine, rather than oviductal, transfer. To date, two approaches to improving horse embryo culture have been explored. The first involved coculture of presumptive zygotes with somatic cells or their products (oviduct epithelial cells [12], cumulus cells [10], equine trophoblast monolayer conditioned medium [13]) and the second involved culturing zygotes in defined cell-free culture media, such as synthetic oviduct fluid (SOF) [14, 15], Dulbecco modified Eagle medium (DMEM) [10], or human tubal fluid [16]. In all of these systems, however, only a low percentage of zygotes developed to the blastocyst stage (average, 15%). Indeed, the fact that temporary transfer of ICSI-derived horse zygotes to the oviduct of progesterone-treated ewes remains the most successful system for producing horse blastocysts ex vivo (rates approach 50% [4]) emphasizes the inadequacy of current IVC systems.

In vitro-produced mammalian embryos tend to exhibit characteristic differences from their in vivo counterparts, including lower cell numbers, an altered inner cell mass (ICM):trophectoderm (TE) ratio, irregularly sized blastomeres, and an increased incidence of cytoplasmic fragmentation, all of which contribute to reduced developmental competence (pig [17], cow [18]). It has also been suggested that apoptosis is a major cause of embryonic arrest in suboptimal culture conditions, such as an excess embryo:medium ratio [19], heat shock [20], excess oxygen free radical concentrations [21], or following exposure of embryos to a high concentration of spermatozoa during IVF [22]. However, apoptosis is also seen in in vivo-produced mammalian blastocysts (mouse [19], humans [23], cattle [24]) and, in these circumstances, has been proposed as a means of eliminating cells that are damaged, nonfunctional, abnormal, or misplaced [23, 25]. Nevertheless, since IVP embryos exhibit relatively high levels of apoptosis (with varying degrees of cytoplasmic fragmentation) and a high incidence of developmental arrest during culture, parameters such as cell number and apoptosis rate may be valuable indicators of the health and developmental capacity of preimplantation embryos.

During IVP of hamster [26], rat [27], and pig [28] embryos, the organization of actin filaments can be affected by environmental conditions such as temperature, pH, and ion concentration. Given that the actin cytoskeleton is critical to the migration of cytoplasmic organelles and the nucleus during oocyte maturation and fertilization and to cell cleavage during mitosis [29], it would not be surprising if culture conditions affected embryo cleavage and development via aberrant cytoskeletal activity. Whether the actin cytoskeleton of IVP horse embryos differs significantly from that of in vivo embryos has yet, however, to be investigated.

An unusual and vital feature of early embryonic development in the horse is the formation of an acellular blastocyst capsule beneath the ZP on Days 6–7 after ovulation, soon after the embryo enters the uterus [30]. After the loss of the ZP, the capsule remains to envelop the conceptus throughout the second and third weeks of pregnancy [31]. Although the precise functions of the capsule are not known [31], it is thought to provide vital mechanical protection during the period when the conceptus is mobile and propelled throughout the uterine lumen by myometrial contractions [32]. The resulting intrauterine migration is essential for the conceptus to distribute its maternal recognition of pregnancy signal to, and thereby inhibit prostaglandin F2{alpha} secretion from, the endometrium sufficiently to prevent luteolysis [33]. Production of the glycoprotein capsule appears to be primarily, or exclusively, a function of trophoblast cells (at least after Day 11 of gestation [34, 35]). However, the presence of large quantities of the endometrial lipocalin, P19 [36], associated with the capsule argues that there may also be a maternal uterine contribution to capsule formation, as does the finding that equine embryos that blastulate in vitro are not able to produce a visible capsule [37, 38]. On the other hand, the suggestion that the ZP is an essential requirement for capsule formation [39] has been disproved by the demonstration that zona-free bisected blastocysts develop an apparently normal capsule after transfer to the uterus of recipient mares [37]. In any case, it has yet to be established whether the initial capsule layer is composed of the same TE-secreted glycoproteins that predominate at later stages and are recognized by the antibody OC-1 [34]. Neither has the influence of IVC on capsule glycoprotein production and coalescence been examined.

The aim of this study was to document the morphological and ultrastructural effects of culturing horse embryos ex vivo in either synthetic oviduct fluid (SOF) [15] or in the oviducts of progesterone-treated ewes. Light and multiphoton scanning confocal microscopy would be used to compare total cell numbers, nuclear morphology, microfilament distribution, and rates of apoptosis in IVP versus in vivo produced embryos. To examine the influence of culture on capsule formation, Days 6–9 in vivo and Days 7 and 10 IVP embryos would be analyzed with respect to capsular glycoprotein expression and distribution.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Unless otherwise indicated, chemicals were purchased from Sigma-Aldrich (Milan, Italy) and plastic ware from Nunc (Roskilde, Denmark).

In Vitro Production of Day 7 Embryos

Collection and culture of cumulus oocyte complexes Ovaries from slaughtered mares were transported to the laboratory in PBS at 25°C. Within 4 h of slaughter, cumulus oocyte complexes (COCs) were recovered by scraping the inside of follicles between 0.5 and 3.0 cm in diameter with a Jacobson curette. The scrapings were washed into Petri dishes using TCM199 supplemented with 25 mM Hepes, 1 mg/ml of BSA, and 10 g/ml of heparin. COCs were classified as either compact (Comp) or expanded (Exp), depending on cumulus and granulosa cell morphology, in the manner described by Hinrichs and Williams [40]. Thus, Comp COCs were those surrounded by a dense cellular mass and with a smooth cumulus hillock and homogenous coloration, whereas Exp COCs were those with cells protruding from the surface of the cumulus hillock and matrix visible between the cumulus cells. COCs were matured by culture for 22–24 h in TCM199 supplemented with 10% fetal calf serum (FCS), 1 µl/ml of ITS (insulin, transferrin, sodium selenite), 1 mM sodium pyruvate, 100 ng/ml of Long-IGF1, 50 ng/ml of Long-EGF, and 0.1 IU/ml each of FSH and LH (Pergovet, Serono, Italy) in four-well plates at 38.5°C in an atmosphere of 5% CO2 in air [15, 41].

Preparation of oocytes and sperm for ICSI After culture, oocytes were separated from their cumulus cells by incubation in 25 µg/ml of hyaluronidase in Hepes-buffered SOF medium [15, 41] and then 2.5 mg/ml of trypsin in PBS for 2 min before aspirating them through a fine pipette. Oocytes with an intact cell membrane were returned to maturation medium for 2–4 h, after which those with an extruded first polar body were selected for sperm injection.

ICSI was performed with frozen/thawed ejaculated sperm from a stallion of proven fertility. One hour before injection, the semen was thawed and the spermatozoa were rinsed free of cryoprotectant by centrifuging them at 750 x g through a discontinuous Percoll density gradient (45%/90%) for 40 min at room temperature. The viable spermatozoa recovered from the bottom of the tube were washed in Ca2+-free TALP [42] and repelleted by centrifugation at 400 x g for 10 min. This second sperm pellet was suspended at a concentration of 4 million sperm/ml in Hepes-buffered SOF medium supplemented with 6 mg/ml of fatty acid-free (FAF) BSA, modified Eagle medium (MEM) amino acids, 1 µg/ml of heparin, 20 µM penicillamine, 1 µM epinephrine, and 10 µM hypothaurine (SOF IVF medium [43]). Just before ICSI, the sperm suspension was diluted 1:1 (vol/vol) with a 12% solution of polyvinylpyrrolidone in SOF IVF medium.

Intracytoplasmic sperm injection Sperm injection was performed as described by Kimura and Yanagimachi [44] using a Piezo micropipette-driving unit (Prima Tech, Tsukuba, Japan) fixed on a micromanipulator (Narishige, Japan) and mounted on an inverted microscope equipped with a 37°C heated stage. A pipette with inner and outer diameters of 50 and 150 µm, respectively, was used to hold oocytes, and a tapered pipette with an inner diameter at the tip of approximately 5 µm was used for sperm injection. A motile sperm was aspirated into the injection pipette and immobilized by applying two or three piezo-pulses to the tail-midpiece junction. The oocyte was held on the holding pipette by suction, with the polar body orientated to the 6- or 12-o'clock position, and the injection pipette was advanced through the ZP at the 3-o'clock position using the piezo-drilling motion. The core of ZP so excised was expelled into the holding medium, and, finally, the injection pipette was advanced through the oolemma using one piezo-pulse and the sperm cell was released into the ooplasm.

Culture of injected oocytes After sperm injection (Day 0), oocytes were cultured in groups of 20 in 20-µl droplets of SOF medium supplemented with MEM amino acids and 16 mg/ml of FAF BSA (SOF-BSA-AA [43]), under mineral oil at 38.5°C in an atmosphere of 5% CO2, 5% O2, and 90% N2. On Day 2 of incubation, the cleavage rate was determined, and morphologically normal two- and four-cell embryos were selected for further culture. The selected embryos were incubated in one of two culture systems, namely, 1) IVC in 20-µl droplets of SOF-BSA-AA, which was partially replaced (by adding 20 µl of fresh medium and then removing 20 µl of the mix) with SOF-BSA-AA on Day 3 and with TCM199 BSA on Day 6, and 2) sheep-oviduct culture after surgical transfer of embryos embedded in agar chips to the ligated oviduct of a ewe implanted with an intravaginal progesterone-releasing device (EAZI-BREED CIDR; InterAg, Hamilton, New Zealand) on the day of transfer, as previously described by Willadsen [45]. For the IVC group, both Exp and Comp COCs were used to produce embryos, whereas for the sheep oviduct group only Comp COCs were used. After an additional 5 days in culture, embryos in the IVC group were assessed, and those lacking cellular compaction and displaying irregular cell sizes were fixed and stained to examine their developmental status. Sheep oviduct embryos were harvested 5 days after transfer into the oviduct, and the rate and quality of Day 7 blastocysts from both systems were scored according to the criteria outlined in the Manual of the International Embryo Transfer Society [46].

Day 10 In Vitro-Produced Embryos

A proportion of the morphologically normal Day 7 blastocysts produced via the sheep oviduct culture system were maintained for a further 3 days in one of two different culture systems: 1) a semidefined culture system consisting of 20-µl droplets of a 1:1 (vol/vol) mixture of DMEM (Gibco BRL, Paisley, Scotland) and TCM199, supplemented with 5% FCS and 5% Serum Replacement (Knockout SR; Gibco BRL) and incubated under mineral oil (the medium was partially replaced on Day 8 of culture), and 2) a cell coculture system consisting of an adult horse skin fibroblast cell (SFC) monolayer and 300 µl of DMEM and TCM199 (1:1) supplemented with 5% FCS and 5% SR in four-well plates. The SFCs were prepared from a small piece of subdermal tissue harvested from the chest of a 5-yr-old Haflinger mare, under local anesthesia. The tissue was sliced finely and digested for 30 min at 38°C in 0.5% (vol/vol) trypsin-EDTA in PBS. The digested fragments were then washed twice with PBS by centrifuging them at 700 x g for 10 min, and the resulting pellet of cells was resuspended in DMEM supplemented with 10% FCS (vol/vol) and cultured at 38.5°C in 5% CO2, 5% O2, and 90% N2. Cells used for coculture had been through one to five passages.

Collection of In Vivo-Produced Horse Embryos

Sixteen early horse conceptuses were recovered from 5–10-yr-old Dutch Warmblood mares by nonsurgical uterine lavage with modified Dulbecco PBS supplemented with 1% FCS, as described by Imel et al. [47]. The conceptuses were recovered 6, 7, or 9 days after ovulation detected during daily ultrasonographic examinations of the ovaries and were thus, respectively, 6.5, 7.5, and 9.5 ± 0.5 days old at the time of collection.

Embryo Evaluation

IVP and in vivo horse embryos of similar developmental stages were compared in two different ways. In experiment 1, Day 7 in vivo embryos and Day 7 IVP embryos from both the IVC and sheep oviduct groups were analyzed for total cell number, apoptotic rate, and microfilament distribution. In experiment 2, Day 10 IVP embryos from both the semidefined and SFC monolayer culture systems were evaluated for total cell number, microfilament organization, and blastocyst capsule formation. Four Day 7 embryos derived from SOF (three) or sheep oviduct culture (one) and a range of Day 6–9 in vivo embryos were also analyzed in this way. In all cases, embryos were measured and assessed morphologically immediately after collection or harvest and before further fixation and labeling.

Morphological assessment Morulae and blastocysts produced in vitro (Days 7 and 10) or in vivo (Days 6, 7, and 9) were measured using a stereomicroscope fitted with an eyepiece micrometer. For Day 10 IVP embryos, separate measurements were made of the part of the embryo that remained enclosed within the ZP and the part that had herniated from the hole made during ICSI. In addition, each embryo was graded morphologically on a scale of 1–4, where 1 represented good quality and 4 was indicative of degeneration. In this respect, in vivo embryos were graded using the system described by McKinnon and Squires [48], whereas an adapted scale based on criteria published in the Manual of the International Embryo Transfer Society was developed for assessing IVP embryos (Table 1; Fig.1). This latter scale included parameters such as degree of compactness, size and appearance of the perivitelline space, color, presence of extruded cells, and the degree of cell granulation and cytoplasmic fragmentation [23]. After morphological assessment, embryos were fixed for 24 h in 4% paraformaldehyde and then stored at 4°C in PBS before staining.


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  		TABLE 1. System used to grade the quality of equine embryos produced in vitro.*



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  		FIG. 1. Photomicrographs to illustrate the quality scoring of Day 7 IVP horse embryos (grades 1–4: good to degenerate). Grading was based on criteria laid down in the Manual of the International Embryo Transfer Society and included degree of compactness, size and appearance of the perivitelline space, color, presence of extruded cells, and the degree of cell granulation and cytoplasmic fragmentation. A and B) grade 1; B and C) grade 2; D) grade 2–3; E) grade 3; F) grade 4; G) grade 4; H) A group of grade 1–2 embryos cultured in SOF medium; In vivo, a grade 2 in vivo early blastocyst for comparative purposes. A–G) Bar = 45 µm; H) bar = 90 µm; and In vivo, bar = 75 µm

Cell number, apoptotic index, and microfilament distribution (experiment 1) Apoptosis was detected using the terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling (TUNEL) technique for DNA fragmentation. First, fixed embryos were incubated twice for 15 min in PBS containing 150 mM glycine and 1 mg/ml of polyvinylalcohol (PVA) to reduce free aldehydes and to block nonspecific reactions. Next, they were permeabilized by immersion for 15 min at 4°C in 0.1% (vol/vol) Triton X-100 in PBS. The permeabilized embryos were then washed twice in PBS containing 1 mg/ml of PVA (PBS-PVA; pH 7.4) before being incubated in 20-µl drops of fluorescein-conjugated dUTP and TdT (TUNEL reagents; Boehringer Mannheim, Roche Diagnostics GmbH, Mannheim, Germany) for 1 h at 37°C in a dark, moist chamber. Following TUNEL, embryos were washed three times in 0.5% (vol/vol) Triton X-100 in PBS containing 1 mg/ml of PVA (PBS-TX100-PVA). To enable microfilament detection, embryos were then incubated for 1 h with 15 IU/ml of AlexaFluor 568 Phalloidin (Molecular Probes Europe BV, Leiden, The Netherlands; A-12380) in PBS-TX100-PVA. Finally, to enable DNA visualization, the embryos were washed twice in PBS-PVA and incubated with 0.1 µg/ml of 4,6-diamino-2-phenyl-indole (DAPI) in PBS for 10 min.

Cell number, microfilament distribution, and capsule immunolocalization (experiment 2) Fixed embryos were washed and permeabilized in 0.1% Triton X-100, as described above. Capsular glycoproteins were labeled using a monoclonal antibody raised against Days 13.5–15.5 equine capsule in a mouse (OC-1) [34]. For the labeling, embryos were first incubated for 45 min in a blocking solution (0.1 M glycine, 1% [vol/vol] goat serum, 0.01% Triton X-100, and 0.5% [wt/vol] BSA) and then exposed to a 1:100 dilution of monoclonal antibody OC-1 in PBS-TX100-PVA for 1 h at 37°C. Next, the embryos were incubated with a 1:300 dilution of a goat anti-mouse IgG coupled to Alexa Fluor 488 (Molecular Probes; A-11029) in PBS-TX100-PVA and then washed twice in PBS-PVA before incubation with AlexaFluor 568 Phalloidin (15 IU/ml in PBS-TX100-PVA) to label the microfilaments. Finally, the embryos were counterstained with DAPI (0.1 µg/ml in PBS) to label the nuclei. As controls, a few embryos were incubated with labeled goat anti-mouse secondary antibody without preexposure to OC-1. In addition, Day 6, 7, and 9 in vivo embryos on which the capsule was clearly identifiable by light microscopy were stained with monoclonal antibody OC-1 as positive controls, whereas in vitro- and in vivo-matured horse oocytes were also stained to control for cross-reactivity of OC-1 with the ZP.

Fluorescence and multiphoton laser scanning microscopy Stained embryos were mounted on glass slides with an antifade (Vectashield, Vector Laboratories, Burlingame, CA). The embryos were examined first using a multiphoton excitation microscopy system combined with a confocal laser scanning microscope (Bio-Rad Radiance 2100 MP) mounted on a Nikon TE300 inverted microscope (Nikon Instruments B.V., Badhoevedorp, The Netherlands). Imaging was performed using a 488-Argon-ion laser and a 543-HeliumNeon laser to simultaneously excite fluorescein and Alexa Fluor 568 (experiment 1) or Alexa Fluor 568 and Alexa Fluor 488 (experiment 2), respectively. DAPI staining was imaged using a 100-fs, pulsed, 780-nm excitation laser source (a mode-locked Titanium-Saphire laser; Tsunami, Spectra Physics, Mountain View, CA). To avoid cross-talk of the images in the photomultiplier channels, specimens were analyzed using a sequential scanning mode. Images were recorded digitally and processed using Adobe Photoshop 5.5 software (Adobe Systems Inc., Mountain View, CA). After multiphoton laser scanning microscopy (MPLSM), mounted embryos were flattened by pressing on the coverslip to enable easier counting of nucleus numbers using a conventional immunofluorescence microscope equipped with an eyepiece counting grid.

The method for calculating apoptosis rates was adapted from Spanos et al. [49]. Stained nuclei were categorized as follows: 1) compact DAPI—interphase nuclei with a distinct round outline, uniform DAPI staining, and no TUNEL (Fig. 2A); 2) mitotic DAPI—nuclei in prophase, metaphase, or anaphase of mitosis with no TUNEL); 3) compact TUNEL—nuclei with strong DAPI and uniform TUNEL); 4) fragmented nuclei with TUNEL; and 5) fragmented nuclei without TUNEL. For each embryo, all nuclei were counted and categorized into one of these five groups. The cells deemed to be apoptotic were all those with TUNEL nuclei plus those with fragmented nuclei but no TUNEL. The following indices were then calculated:

a. apoptotic index = apoptotic nuclei/total nuclei x 100;
b. fragmented nucleus index = fragmented nuclei/total nuclei x 100;
c. mitotic index = nuclei in mitosis/total nuclei x 100.



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  		FIG. 2. The effect of cumulus morphology at the onset of IVM (Comp or Exp) and subsequent culture system (SOF medium or sheep oviduct) on the mean (+SEM) percentages of cells showing evidence of apoptosis, nuclear fragmentation, or cell division (mitosis) in Day 7 IVP horse embryos. Embryos derived from Comp COCs and cultured in SOF had a significantly higher proportion of apoptotic and fragmented nuclei. Within a group, different superscripts denote values that differ significantly (P < 0.05)

Statistical Analysis

All culture experiments included 5–7 replicates, and statistical analyses were performed using GraphPad Prism (San Diego, CA). Cleavage and in vitro embryo development rates were compared using the Fisher exact contingency test. The effect of culture condition on total cell numbers and on the apoptotic, fragmentation, and mitotic indices was examined using either unpaired Student t-tests or a one-way ANOVA followed by pairwise multiple comparisons (Bonferroni t-test), after testing for normality (Kolmogorov-Smirnov test) and equivalence of variances (Levene Median test with Bartlett test correction). Differences were considered statistically significant if P < 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
In Vitro Embryo Development

A total of 666 oocytes that displayed a first polar body after 24–26 h of IVM were subjected to ICSI. Of these, 349 and 317 were derived from COCs categorized as Comp or Exp at the onset of IVM, respectively. There was, however, no effect of cumulus morphology at the onset of IVM on the cleavage rate after ICSI (61.3% versus 63.4% for Comp and Exp COCs, respectively; Table 2). On the other hand, there was a significant effect of culture system on the embryo development rate, since zygotes from Comp COCs incubated in the sheep oviduct system yielded a significantly higher blastocyst rate (16% of injected and 23.7% of cleaved oocytes) than SOF IVC with either Comp (6.3% and 12.6%) or Exp (9.4% and 15%) COCs (Table 2).


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  		TABLE 2. Cleavage and development of horse embryos produced by ICSI of IVM oocytes, which had Comp or Exp COCs at the onset of IVM, and subsequent culture for 7 days in SOF medium or the oviduct of a progesterone-treated ewe.*

Morphology and cellular characteristics of Day 7 embryos (experiment 1) Of the 69 Day 7 embryos produced by ICSI and either culture in SOF (n = 41) or temporary transfer to the sheep oviduct (n = 28), 47 were analyzed with respect to cell number, apoptosis, and microfilament organization. These were compared with 10 in vivo produced Day 7 embryos.

Embryo diameter and total cell number Day 7 in vivo embryos were significantly bigger than Day 7 IVP embryos (mean ± SEM embryo diameters, 374 ± 64.1 mm, 139.9 ± 1.6 mm, and 138.1 ± 2.5 mm for in vivo, SOF, and sheep oviduct embryos, respectively; Table 3). Not surprisingly, in vivo embryos also contained significantly more cells (1736 ± 568; range, 176–5720) than SOF (116 ± 15; range, 20–363) or sheep oviduct (86 ± 9; range, 31–141) produced embryos. With regard to the IVP embryos, COC morphology at the onset of culture did not affect eventual blastocyst diameter but did significantly influence Day 7 cell number, which was significantly higher for Exp COCs (133 ± 16) than for Comp COCs subsequently cultured via either the SOF or sheep oviduct systems (83 ± 31 and 86 ± 9, respectively; Table 3).


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  		TABLE 3. Diameter and total cell number of Day 7 horse embryos produced in vivo or in vitro following ICSI of oocytes from Comp and Exp COCs and culture in SOF (IVC) or from compact COCs and temporary transfer to the oviduct of a sheep.*

Apoptosis and nuclear morphology Apoptosis occurred at a very low rate in in vivo embryos, with only 4 of the 10 embryos containing apoptotic cells and only 0.5% of their nuclei being classified as apoptotic. By contrast, all IVP embryos contained apoptotic nuclei and apoptosis rates were much higher. For this reason, the apoptotic, mitotic, and fragmentation indices were compared between the different groups of IVP embryos. In this respect, the proportion of apoptotic cells was significantly higher in embryos produced from Comp COCs and SOF (21.5% ± 6.4%; Fig. 2) than Exp COCs and SOF (9.6% ± 1.3%) or Comp COCs and sheep oviduct transfer (6.5% ± 1.5%). A similar pattern was seen for the incidence of fragmented nuclei (13.3% ± 3.5% vs. 4.9% ± 0.9% and 4.5% ± 1% for Comp COCs SOF, Exp COCs SOF, and Comp COCs sheep oviduct, respectively), but there was no apparent effect of COC morphology or culture system on the mitotic index. More detailed analysis of the different classes of apoptosis (condensed TUNEL positive, fragmented TUNEL positive, or fragmented TUNEL negative) demonstrated that the bulk of the between-IVP group difference was in the proportion of fragmented TUNEL-positive cells; indeed, the proportion of TUNEL-negative apoptotic cells did not differ between groups (Fig. 3). When apoptosis rates were compared between IVP embryos of different quality, it was found that embryos considered transferable (grades I and II; 76% of all IVP embryos) had lower apoptosis rates (9% of cells) than embryos not considered fit for transfer (grades III and IV; 22% apoptotic cells).



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  		FIG. 3. The effect of cumulus morphology at the onset of IVM (Comp or Exp) and culture system (SOF or sheep oviduct) on the rate of different categories of apoptosis in Day 7 horse embryos (mean ± SEM). Day 7 embryos derived from Comp COCs and cultured in SOF had a significantly higher proportion of fragmented TUNEL-positive nuclei. Within a group, different superscripts denote values that differ significantly (P < 0.05)

Microfilament organization Representative MPLSM photomicrographs of Day 7 in vivo and in vitro embryos are shown in Figure 4. IVP embryos were not only smaller and had fewer cells than their in vivo counterparts, they were also more compact, such that most were categorized as morulae (Fig. 4, A and B) or early blastocysts (Fig. 4, C and D), whereas in vivo Day 7 embryos were mostly expanded blastocysts (Fig. 4, H and I). The distinction between ICM and TE was thus more difficult to discern in Day 7 IVP embryos, which did not have the clear nucleus-dense domain seen in in vivo embryos. Apoptotic cells were scattered and not concentrated in particular areas but were seen in higher numbers in poor-quality IVP embryos (Fig. 4, E and F). Microfilaments were primarily located along the inside of the cell plasma membranes, adjacent to the cell borders. In in vivo blastocysts, microfilament labeling of the contiguous cell borders was intense and homogeneous throughout the whole embryo. By contrast, in IVP embryos microfilament labeling was generally less intense and tended to be more marked around the periphery of the embryo (Fig. 4, A and D) and weaker within, with some areas almost entirely lacking labeling (Fig. 4, G). On some occasions, the actin labeling in IVP embryos appeared to have agglomerated at the junctions of several cells (Fig. 4, E). The blastocoele cavity of IVP blastocysts tended to be small and irregular with its borders poorly stained for microfilament (Fig. 4, C), especially when compared with the large, well-delineated cavity of in vivo blastocysts (Fig. 4, H).



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  		FIG. 4. Confocal laser scanning photomicrographs to demonstrate the features of and differences between morphology and microfilament distribution in in vivo and IVP horse embryos. In all cases, nuclei are colored blue, microfilaments red, and TUNEL nuclei green (these nuclei contain DNA breaks and are considered to be apoptotic). IVP embryos had relatively few cells and were relatively compact, such that most were categorized as morulae (A and B) or early blastocysts (C and D). Day 7 in vivo embryos were mostly expanded blastocysts (H and I) with a clear distinction between TE and ICM. Apoptotic cells tended to be scattered but more numerous in poor-quality IVP embryos (E and F; f denotes a fragmented nucleus). Microfilaments lined the inside of the cell plasma membrane and were intense and sharp in in vivo embryos. In IVP embryos, microfilament labeling was generally less intense but tended to be stronger around the periphery of the embryo (A and D) and weaker to almost absent in the center (G). On some occasions, the actin filaments were clumped at the junction of several cells (E). The blastocoele cavity of IVP blastocysts tended to be small and much less clearly delineated by a microfilament border (C) than that of in vivo blastocysts (H). A–G) Bar = 30 µm; H) bar = 40 µm; and I) bar = 120 µm. Cp, compact DAPI-labeled nucleus; f and c, fragmented and compact TUNEL nuclei, respectively

Capsular glycoprotein expression (experiment 2) In total, 20 Day 10 IVP embryos were stained with monoclonal antibody OC-1 (Table 4) to examine capsule formation. These embryos had been produced by 5-day culture in sheep oviducts followed by 3 days either in a cell-free system (n = 10) or with a monolayer of adult horse fibroblast cells (n = 10). All Day 10 IVP embryos were classified as partially hatched, since part of the embryo had herniated via the hole made in the ZP during ICSI. In these embryos, a distinct layer of OC-1-positive capsule was visible lining the inside of the ZP (Fig. 5, A and B) and extending into the transzonal channels of the latter (Fig. 5, E). When the ZP was separated from its contained embryo by micromanipulation, the capsular material remained stuck to the ZP and not to the TE (Fig. 5, F), demonstrating that it was more intimately associated to the former. In the area of embryo herniated from the ZP, capsular glycoprotein was present on the apical surface of the TE cells (Fig. 5, A and B) as scattered small patches that were not assembled into a confluent layer (Fig. 5, I and H). Day 7 IVP embryos (three from SOF and one from sheep oviduct cultures) showed only weak OC-1 labeling in scattered patches on the apical surface of TE cells, again without assembly into a confluent capsule and, in these cases, with little infiltration into the substance of the ZP (Fig. 5, C and D). By contrast, in two Day 6 in vivo embryos examined (categorized as late morulae), a clear, thick confluent capsule was sandwiched between the relatively thick ZP and the trophectodermal surface, and there was no infiltration of glycoproteins into the transzonal channels of the ZP (Fig. 5, J). Older in vivo hatched and expanded blastocysts had a complete capsule apposed tightly to the trophectodermal surface, and these capsules displayed the classic bilaminar appearance (Fig. 5, K) described previously by Oriol et al. [34].


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  		TABLE 4. Immunolocalization of capsular material on early in vivo and IVP horse embryos.*



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  		FIG. 5. Confocal laser scanning photomicrographs to show the distribution of OC-1-reactive molecules (capsular glycoproteins) in IVM oocytes and IVP and in vivo embryos. In all cases, nuclei are labeled blue, microfilaments red, and capsular glycoproteins green. A) Capsule material lining the inside of and permeating through the ZP of a Day 10 IVP embryo. OC-1 staining is also seen (arrow) on the outer surface of the TE cells that have herniated from the ZP via the hole made during ICSI. B) Day 10 IVP embryo with strong OC-1 staining on the herniated ZP-free TE cells. Cell density in the ZP-enclosed portion of the embryo is much higher than in the "unrestricted" herniated area. C) Capsular material on the trophectodermal surface of a Day 7 embryo derived from sheep oviduct culture. D) Accumulation of capsule material beneath the ZP of a Day 7 embryo derived from IVC. E) Capsular material permeating through the transzonal channels of a Day 10 IVP embryo. F) Separation of the ZP from the TE by micromanipulation demonstrated that the pseudocapsule was adhered to the ZP rather than the TE. G) Day 10 IVP embryo with a dense actin "neck" between the zona-contained and herniated parts of the embryo. H) Herniated part of a Day 10 IVP embryo with considerable quantities of capsular material on the surface but without coalescence into a confluent layer. I) A Day 10 IVP embryo that, during culture with a fibroblast monolayer, attached to the surface of the well. It is presumed that failure of OC-1-reactive glycoproteins to coalesce prevented them from exerting the antiadhesive properties that have been ascribed to the capsule. J) Day 6.5 in vivo early blastocyst with a distinct, well-defined capsule and a very clear microfilament cytoskeleton delineating individual cells. K) High magnification image of the capsule and TE of a Day 7 in vivo blastocyst to show the bilaminar appearance of the capsule after staining with monocloal antibody OC-1. L) COC stained after IVM to demonstrate that monoclonal antibody OC-1 does not cross-react with ZP. A–C, I, and K) Bar = 50 µm; D–H, J, and L) bar = 10 µm

Because OC-1 showed such strong affinity for the ZP of IVP embryos, a number of in vitro matured oocytes (n = 4) were also stained with OC-1 to ensure that there was no cross-reaction with ZP after IVC. In these cases, there was no OC-1 staining of any part of the ZP (Fig. 5, L), demonstrating clearly that staining of the ZP in IVP embryos was a function of embryonic secretion of OC-1 reactive glycoproteins.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
This study demonstrated that IVP horse embryos differ markedly from their in vivo-produced counterparts by having fewer cells, lower morphological quality scores, and a higher incidence of apoptotic cells (Table 3). These observations are similar to those for IVP bovine embryos, which are consistently of lower quality than embryos produced in vivo, both in terms of morphological quality [50] and with regard to the expression of developmentally important genes [51]. In cattle, these differences have been shown to adversely affect embryo survival following cryopreservation and to compromise embryonic development [52]. On the other hand, because only a few foals have so far been produced from IVP embryos [4, 10, 11], it is too early to determine if and to what degree in vitro production adversely affects embryo development in this species. It is, however, clear that despite the apparently poor quality and low cell numbers of IVP equine embryos, they are able to establish pregnancies following transfer to recipient mares [10, 11].

As in previous studies [4, 41], embryo development was more efficient when zygotes were transferred temporarily to the oviduct of progesterone-treated sheep (23% blastocysts) than when they were cultured exclusively in vitro (14%). However, the number of cells in Day 7 IVP embryos derived from Comp COCs and either cultured in SOF medium or transferred to a sheep's oviduct did not differ. By contrast, cell numbers were higher in Day 7 IVP embryos originating from Exp COCs and cultured in SOF medium. The effects of this relatively rapid increase in cell number in vitro, of embryos derived from Exp compared with Comp COCs, on later embryo development is however difficult to predict. At first glance, a higher cell number would appear to be a positive sign of embryo vitality. On the other hand, in other species, relatively rapid development in vitro has proven detrimental to the resulting offspring [53]. That the rate of apoptosis (cells with DNA strand breaks and/or nuclear fragmentation) was significantly higher in SOF than sheep oviduct embryos (22% vs. 7%) and much higher in both than in in vivo-produced embryos suggests a detrimental effect of culture on embryo development. Similarly increased rates of apoptosis have been observed in cattle [24] and pig [54] embryos produced in vitro, and it has been suggested that suboptimal culture conditions may cause the proportion of affected cells to increase to a level where further embryonic development is critically compromised, particularly in embryos with low cell numbers [19, 55]. On the other hand, apoptosis has also been proposed to be a physiological process during mammalian preimplantation embryogenesis, acting as a quality control mechanism to eliminate aberrant cells [22, 25]. In either case, the marked difference in the proportion of apoptotic cells between in vivo and IVP horse embryos suggests that culture adversely affects embryo development.

IVP embryos were produced at similar rates from oocytes that had a Comp or an Exp cumulus at the onset of IVM, although the overall percentage of injected oocytes developing into blastocysts was low (mean, 14%). Previous studies have suggested a greater ability of oocytes with an expanded cumulus at recovery to form a male pronucleus after ICSI [16, 56], although no differences in nuclear maturation rates were observed. One possible explanation for this finding is that COCs with an expanded cumulus had better cytoplasmic maturation and were thereby better able to support embryo development. Although the current study did not show any significant difference in cleavage rates between Exp and Comp COC oocytes, embryos resulting from SOF culture after ICSI of Exp COC oocytes had more cells and a lower incidence of apoptosis than those from Comp COCs. Interestingly, the major difference in the proportion of apoptotic cells was a higher level of cells with fragmented nuclei in Comp COC derived embryos. Apoptosis is considered to progress from condensation of chromatin and cytoplasm to DNA fragmentation accompanied by indentation and convolution of the nuclear membrane. TUNEL labels the DNA breaks in situ and thus identifies cells fairly early in the process of apoptosis [57], whereas fragmentation of the convoluted nucleus with blebbing and fragmentation of the whole cell into membrane-bound apoptotic bodies occurs much later in the cascade (for review see Wyllie et al. [58]). The raised incidence of nuclear fragmentation in Comp COC-derived embryos is therefore indicative of advanced apoptotic changes, which could derive from a higher sensitivity to culture conditions and/or presence of more serious defects in these embryos [22]. Since the quality (based on apoptotic index) of embryos derived from Comp COCs was significantly improved when zygotes were transferred to and cultured in sheep oviducts, sensitivity to the culture conditions is the most likely critical factor. Overall, these data suggest that current IVM protocols are suboptimal with respect to the acquisition of developmental competence by equine oocytes, especially for those less advanced oocytes with a compact cumulus at the time of recovery.

Staining of the microfilament cytoskeleton helped to identify differences in cellular organization between in vivo and in vitro embryos. In vivo Day 7 embryos were expanded blastocysts in which the microfilaments delineated the cell borders and highlighted the division between a distinct ICM and a TE layer (Fig. 4, F). By contrast, IVP embryos were compact without or with only a small blastocoele cavity and patchy microfilament distribution. Actin microfilaments play an important role in cell cleavage during embryo development [59] and are essential for the distribution of mRNA and organelles, such as mitochondria and the Golgi apparatus, between daughter cells [60, 61]. The polymerization and depolymerization of actin filaments are, however, dynamic processes susceptible to disruption by environmental conditions (such as temperature, pH, culture medium [26]). Moreover, microfilament disruption can adversely affect the structural integrity of cells, with serious consequences for their metabolic activity. Abnormal actin filament distribution has been suggested as a reason for abnormal embryo cleavage in IVP pig embryos [28], because embryo division can be similarly blocked by cytochalasin D, an inhibitor of microfilament polymerization. In general, cells store a large pool of nonfilamentous actin (G-actin [62]) to maintain their ability to quickly reorganize filamentous actin in response to environmental changes or need. Whether the pool of G-actin is low or impaired in IVP horse embryos, thereby decreasing their potential to overcome suboptimal culture conditions, has yet to be investigated.

The combination of monoclonal antibody OC-1 and confocal microscopy enabled three-dimensional imaging of capsular glycoprotein distribution in early embryos. In turn, this enabled us to demonstrate that the capsule assumes its classic bilaminar appearance around in vivo embryos soon after its initial formation (Fig. 5, K); previously, OC-1 expression and the appearance of the capsule after OC-1 staining had not been reported for embryos recovered earlier than Day 11 of gestation [34]. Production of capsular glycoproteins by the trophoblast cells of IVP embryos was also demonstrated by OC-1 labeling of the apical surface of these cells, thereby confirming the hypotheses that the initial layer of capsule is formed from OC-1-reactive glycoproteins secreted by early TE cells independent of a maternal (endometrial) input [35]. Intriguingly, in vitro, the capsular glycoproteins failed to assemble into a normal and complete capsule enveloping the embryo, suggesting either that some aspect of the uterine environment is necessary for glycoprotein coalescence or that some aspect of the in vitro environment otherwise hindered capsule formation. In the former respect, because the glycoproteins of the capsule are mucinlike, Oriol et al. [34] postulated that they, like other mucins, may coalesce to form a gellike mucous layer by hydration and cross-linking [63] after they have accumulated in sufficient quantities on the surface of the TE. Oriol et al. [34] further suggested that failure of capsule formation in vitro might thus be due to dispersal of the glycoproteins into the culture medium or unsuitability of the microenvironment for hydration and cross-linking. The current study suggests that the former is a minor problem, since capsule glycoproteins were found on the surface of the trophoblast and lining and permeating into the ZP. Nevertheless, it cannot be discounted that a degree of dispersal prevented glycoprotein concentrations reaching a critical concentration needed to initiate coalescence. In this latter respect, it is likely that the absolute production of capsular glycoproteins by IVP embryos was relatively low, due to their low cell numbers, and it is also possible that the ICSI-derived hole in the ZP would have hindered the accumulation of the glycoproteins. In addition, there are comparable reports of reduced secretion of other high-molecular-weight mucins in culture, such that they fail to assemble into a mucous layer [64].

In the present study, considerable quantities of capsular material were detected as an accumulation between the trophoblast and the ZP of IVP embryos (Fig. 5, D). This was reminiscent of the flocculent material detected between the trophoblast and ZP of in vivo embryos by Flood et al. [30] and Wilson et al. [65], which they postulated, but could not prove, to be capsule precursor material. A further striking feature of IVP embryos was the level to which the ZP was lined and infiltrated with capsular glycoproteins, something not seen in in vivo embryos. The possibility that monoclonal antibody OC-1 had cross-reacted with the ZP per se was ruled out by the failure to label the ZP of either in vivo embryos or IVM oocytes (Fig. 5, L). Instead, it appears that during culture in vitro capsule material adheres to the inside of the ZP and permeates through the cumulus cell-created transzonal channels, presumably as a consequence of the failure of the glycoproteins to coalesce on the trophectodermal surface. By contrast, in vivo capsule formation does not require the presence of a ZP, since zona-free bisected blastocysts have been shown to form a capsule after transfer to the uterus of recipient mares [37]. This again indicates that the greatest obstacle to capsule formation in vitro is the failure of the microenvironment to replicate the conditions necessary for glycoprotein cross-linkage. In Day 10 IVP embryos, a pseudocapsule formed as an inner lining to the ZP (Fig. 5, E) that was not present around ZP-free areas of embryo (Fig. 5, H). This capsule was tightly adhered to the ZP, as demonstrated during removal of the latter by micromanipulation (Fig. 5, F); the capsule peeled away with the ZP.

In the current study, in vitro cultured embryos escaped from their ZP by herniating through a hole in the ZP (in this case the hole made during ICSI), as previously reported by Hinrichs et al. [38] and Hochi et al. [66]. In other species, the exact mechanism of hatching from the ZP is also unclear, although it is thought that, in vivo, proteolytic enzymes released by the maternal endometrium are most likely to be responsible for ZP dissolution [67]. The current observations reiterate that loss of the horse ZP in vitro is different from the apparently rapid loss in vivo (intermediate stages are rarely found), during which the ZP is thought to be attenuated and ruptured by a combination of a uterine zonalytic and blastocyst expansion [30, 68]. Although the uterus seems thus to critically influence the mode of zona loss from horse embryos, the existence of a uterine zonalytic has yet to be demonstrated in this species (there appears to be no uterine zonalytic in cattle; [69]). It is therefore tempting to speculate that the essential roles of the uterus during physiological zona loss and capsule formation may be linked. Finally, it has been proposed that the capsule is essential to the survival of horse embryos in vivo because embryos transferred after capsule removal either do not develop into pregnancies [70] or do so only after forming a new capsule [37]. If IVP embryos do not form a normal capsule during culture, then it is likely that transfer to the uterus of the recipient mare at a stage when capsule formation can still occur will be essential to the success of this procedure. Since the transfer of Day 7 IVP late morulae or early blastocysts has resulted in pregnancy in mares, it must be assumed that subsequent capsule coalescence can and does occur.

In summary, the present study represents the first detailed description and comparison of the morphological, cytoskeletal, and developmental characteristics of in vitro- and in vivo-produced horse embryos. Day 7 IVP embryos were smaller, had fewer cells, and were more compact than in vivo embryos of similar age. In addition, Day 7 IVP embryos had a small or nonexistent blastocoele cavity and an indistinct ICM and had still not properly (or normally) hatched after 10 days of culture. IVP embryos also displayed high percentages of apoptotic cells (10% compared with 0.3% for in vivo embryos), a disturbed pattern of microfilament distribution, and irregularities in cell size and shape. Finally, although IVP embryos remained viable, continued to develop for at least 10 days in vitro, and secreted capsular glycoproteins, the latter failed to coalesce to form a confluent capsule, a structure that is a prominent and apparently vital feature of in vivo blastulation.


   ACKNOWLEDGMENTS
 
The authors would like to thank Professor K.J. Betteridge for the generous gift of the monoclonal antibody OC-1 and for his helpful comments on the manuscript. We would also like to thank the research team at the Laboratorio di tecnologie della Ripdroduzione (Cremona, Italy) for their assistance in the production of IVP embryos. In particular, we thank Drs. N. Ponderato, S. Colleoni, and G. Crotti for their excellent technical support and Dr. R. Duchi for performing the sheep oviduct transfers. From Utrecht University, we thank T. Tharasanit for providing the in vivo embryos and Dr. A. de Graaf of the Centre for Cell Imaging at the Department of Biochemistry and Cell Biology for assistance with MPLSM. The MPLSM was purchased with funding from the Dutch Council for Scientific Research, Division of Medical Sciences (NW-MW).


   FOOTNOTES
 
1 Correspondence: Jordi L. Tremoleda, Department of Equine Sciences, Section of Reproduction, Faculty of Veterinary Medicine, Utrecht University, Yalelaan 12, 3584 CM Utrecht, The Netherlands. FAX: 31 30 2534811; J.Tremoleda{at}vet.uu.nl Back

2 Deceased Back

Received: 23 April 2003.

First decision: 16 May 2003.

Accepted: 29 July 2003.


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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