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Degeneration of cryopreserved bovine oocytes via apoptosis during subsequent culture
Cryobiology 47 (2003) 73–81 www.elsevier.com/locate/ycryo
Degeneration of cryopreserved bovine oocytes via apoptosis during subsequent cultureq Hongsheng Men,*,1 Rick L. Monson, John J. Parrish, and Jack J. Rutledge Department of Animal Sciences, University of Wisconsin-Madison, 1675 Observatory Drive, Madison, WI 53706, USA Received 21 February 2003; accepted 4 July 2003
Abstract Cryopreservation causes a significant proportion of bovine oocytes to undergo degeneration during subsequent culture. We investigated the degeneration mechanism of cryopreserved oocytes. In vitro matured bovine oocytes were vitrified by the open-pulled straw (OPS) method. In each replicate, a group of oocytes were randomly taken after warming to determine oocyte survival by both morphological evaluation and propidium iodide vital staining. The remainders were evaluated by morphological criterion. Morphologically intact oocytes were co-incubated with frozenthawed spermatozoa for subsequent development. In situ examination of DNA breaks in oocytes and embryos was conducted using a Fluorescein-FragEL DNA fragmentation detection kit. A caspase-3 detection kit was used to detect caspase-3 activity in oocytes and embryos. Most of the oocytes survived cooling and warming processes as assessed by both morphological evaluation and vital stain. During subsequent culture, some degenerating oocytes displayed observable apoptotic morphology, such as cytoplasmic condensation, cytoplasmic fragmentation, and formation of apoptotic bodies. Biochemical markers of apoptosis, such as apoptotic DNA fragmentation and activation of caspases, were detected not only in oocytes having typical apoptotic morphology, but also in oocytes without observable apoptotic morphology. In embryos, positive signals for both biochemical markers were detected in blastomeres. This experiment suggests that cryopreserved bovine oocytes degenerate via apoptosis during subsequent culture. Ó 2003 Elsevier Inc. All rights reserved. Keywords: Apoptotic cell death; Caspase-3 activity; DNA fragmentation; Vitrification
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This work was supported by the Babcock Institute for International Dairy Research and Development, University of Wisconsin-Madison, and by a grant from USDA/CSREES 2001-52 101-11318. * Corresponding author. Fax: 1-573-8847521. E-mail address: [email protected] (H. Men). 1 Present address: Department of Veterinary Pathobiology, University of Missouri-Columbia, 1600 E. Rollins Street, Columbia, MO 65211, USA.
A dominant feature of mammalian (especially human and domestic animal) oocyte cryopreservation by current protocols is loss of developmental competence. Cryopreservation causes a significant increase in the rate of degeneration in oocytes and embryos derived from cryopreserved oocytes during culture [18,19]. Current efforts in the improvement of oocyte cryopreservation concentrate on increasing the cooling rate to minimize cryoinjuries [20,38]. It may be possible to improve
0011-2240/$ - see front matter Ó 2003 Elsevier Inc. All rights reserved. doi:10.1016/S0011-2240(03)00070-1
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oocyte cryopreservation by inhibiting the initiation of oocyte degeneration during subsequent culture if the degeneration mechanism of cryopreserved oocytes can be characterized. Cells normally degenerate by one of two mechanisms: apoptosis and necrosis. Apoptosis is an endogenous cell degeneration mechanism that is necessary for normal development and tissue homeostasis [43]. The apoptotic process can also be activated in response to a wide variety of nonphysiological stimuli, such as temperature, toxicants, oxidative stress, and X-radiation [41]. Morphological characteristics of apoptotic cells are cell blebbing, margination of the chromatin, and the formation of apoptotic bodies (membranebounded cytoplasmic bodies) [42]. These morphological changes are a result of the activation of a group of proteases, such as caspase-1, caspase-2, caspase-3, etc. Caspases exist in cells as inactive proenzymes. Upon apoptosis initiation, they are activated by cleavage at specific aspartate residues [6]. A variety of proteins, such as PARP, histone H1, DNA-dependent protein kinase, and lamins are destroyed by the action of caspases and result in the morphological changes in apoptotic cells. Generation of DNA fragments of 180–200 bp, a characteristic of apoptosis, is a result of caspaseactivated endonucleases [23,42,43]. On the other hand, when cells experience a wide departure from physiological conditions, such as injury or trauma, another mechanism, necrosis, takes over as the mechanism for cell degeneration. Necrotic cell death is characterized by cell distension followed by lysis and leakage of intracellular contents. Nuclear DNA is randomly cleaved and forms a ‘‘smear’’ in agarose gel after electrophoresis [1,41]. During germ cell development, apoptosis also plays a significant role in germ cell homeostasis. Apoptotic cell death can be observed as early as the primordial germ cell proliferation stage [40] and continues throughout spermatogenesis [15,30] and oogenesis [31,36]. Up to 75% of male germ cells are lost during spermatogenesis [30]. During ovarian development, over two-thirds of oocytes are eliminated through apoptosis before birth and a large number of remaining oocytes are lost through apoptosis during follicular atresia [36].
Ovulated oocytes also undergo degeneration if fertilization does not occur. It is generally thought that apoptosis is the degeneration mechanism of ovulated oocytes [9,28,34] although Van Blerkom and Davis [39] present evidence otherwise. In vitro cultured mammalian (including bovine) oocytes and embryos without cryopreservation are also degenerated through an apoptotic pathway [10,12,14,26,44]. However, the investigation into the mechanism behind degeneration of cryopreserved oocytes is lacking. It is essential to elucidate the mechanism behind oocyte degeneration after cryopreservation, because cryopreserved oocytes are currently fertilized and cultured to the blastocyst stage in vitro using the same culture procedures developed for non-cryopreserved oocytes. However, there are reasons that cryopreserved oocytes may need a specialized culture system. During cryopreservation, oocytes experience severe adverse physiological conditions. These adverse conditions render oocytes with various cytological injuries [3,32]. In addition to cytological abnormalities, cryopreservation may also result in biochemical abnormalities to oocytes as it has to other cell types. For example, in bone marrow cells, cryopreservation can activate intracellular proteases and the activated proteases in turn cause artificial cleavage of apoptosis-related proteins [33]. Therefore, prevention of the initiation of apoptosis would in turn increase survival of cryopreserved cells. Many cell types are known to require modification of freezing solutions or culture systems for optimal growth or survival. Apoptotic inhibitors in the cryopreservation solution or cold storage solution have enhanced the survival of several cell lines or tissue [2,16,33]. Vascular endothelial growth factor shows a protective effect on cold preserved sinusoidal endothelial cells during storage and culture [25]. Therefore, it would be advantageous in guiding the cryopreservation or the design of culture system for cryopreserved oocytes if we can characterize the degeneration mechanism of cryopreserved oocytes and embryos derived from cryopreserved oocytes. Herein, we test the hypothesis that cryopreserved oocytes and embryos derived from cryopreserved oocytes degenerate via apoptosis.
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Materials and methods Unless otherwise indicated, all chemicals were purchased from Sigma Chemical (St. Louis, MO). Maturation, fertilization, and culture of bovine oocytes In this study, we employed a bovine oocyte maturation, fertilization, and culture system that could consistently give 80% cleavage rate and 30% blastocyst formation rate (out of cleaved embryos). Cumulus–oocyte complexes were obtained by aspiration of 3–8 mm follicles from ovaries collected from a local slaughterhouse. Oocytes with uniformly granulated cytoplasm and at least one layer of cumulus cell investment were washed three times in TL–Hepes (BioWhittaker, Walkersville, MD) and matured for 22 h at 38.5 °C, 5% CO2 in air with maximal humidity in 50 ll drops of medium TCM-199 (BioWhittaker) with the following supplementations: 10% FBS (Gibco-BRL, Grand Island, NY), 0.2 mM Na-pyruvate, 3 lg/ml FSH (National Institutes of Health, Bethesda, MD), 3 lg/ml LH (National Institutes of Health), 1 lg/ml estradiol, and 25 lg/ml gentamycin. Oocytes were fertilized with 45–90% Percoll gradient selected living sperm from frozen semen at a final concentration of 1 106 sperm/ml in a 50 ll fertilization medium drop under 5% CO2 in air with maximal humidity at 38.5 °C [27]. The fertilization medium used is a modified TyrodeÕs medium supplemented with 0.2 mM Na-pyruvate, 6 mg/ml fatty acid free BSA, 2 lg/ml heparin, 20 lM penicillamine, 10 lM hypotaurine, and 1 lM/ml epinephrine. After 22 h of fertilization, putative zygotes were cultured in 50 ll drops of KSOM (Specialty Media, MD) with 3 mg/ml BSA as the protein source. Vitrification of bovine oocytes Protocols of oocyte vitrification with openpulled straws (OPS) were as described by Vajta et al. [38]. After partial removal of cumulus cells, mature oocytes were first dehydrated by 45 s exposure to 10% (v/v) ethylene glycol (EG) + 10% (v/v) dimethyl sulfoxide (Me2 SO) + TL–Hepes with
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20% (v/v) FBS supplementation and then in vitrification solution (20% EG + 20% Me2 SO + 0.5 M sucrose in TL–Hepes + 20% FBS). Oocytes with minimal amount (1–2 ll) of vitrification solution were picked up in the narrow end of the OPS by capillary action. The straw was plunged directly into liquid nitrogen within 30 s of contacting the concentrated vitrification solution. Vitrified oocytes were stored in liquid nitrogen for 1 or 2 days. Warming was conducted by immersing the narrow end of the OPS into TL–Hepes with 0.25 M sucrose and 20% FBS for 1 min at 37 °C. Rehydration was conducted by subsequent washes for 5 min in each wash: once in TL–Hepes with 0.15 M sucrose + 20% FBS and then twice in TL–Hepes + 20% FBS. Viability assay Viability of warmed oocytes was determined based on visual examination of the integrity of the oocyte membrane, zona pellucida, and the normality of the cytoplasm immediately after warming. The validity of morphological classification was confirmed by vital staining with propidium iodide. To perform vital stain, samples of visually viable and dead oocytes were mounted separately on microscope slides and then covered by coverslips with the support of Vaseline–wax mix. Oocytes were stained with propidium iodide at a final concentration of 0.2 mg/ml in PBS. The oocytes were examined under a Nikon Diaphot microscope equipped with epifluorescent microscopy. A combination of 510–560 nm excitation filter and 590 nm barrier filter was used to examine PI fluorescence. The remaining oocytes were evaluated by morphological observation. Visually dead oocytes were discarded, and the morphologically intact oocytes were subject to fertilization and culture. Morphological characterization of apoptosis in oocytes Morphological characterization of apoptosis was carried out under a Bausch and Lomb stereomicroscope. Oocytes having the observable features of apoptotic morphology, such as condensed cytoplasm, fragmented cytoplasm and
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apoptotic bodies were considered to undergo apoptotic cell death. Photographs were taken with a Zeiss inverted microscope equipped with differential interference contrast optics using ASA 400 Fuji black and white film with 200 magnification. Detection of DNA fragmentation in bovine oocytes and embryos DNA integrity of degenerating oocytes or embryos with fragmented blastomeres was evaluated by using DNA fragmentation detection kit (Fluorescein-FragEL, Oncogene Research Products, Boston, MA). The principle of Fluorescein-FragEL is that terminal deoxynucleotidyl transferase (TdT) catalyzes the addition of fluorescein-labeled and unlabeled deoxynucleotides to the 30 -OH ends generated by endonucleases during apoptotic degeneration. Oocytes or embryos were first fixed in 4% paraformaldehyde in PBS + 1% BSA for 15 min and permeabilized for 5 min in 0.01% Triton X-100 in PBS–BSA at room temperature. After three washes in PBS–BSA, oocytes/embryos were incubated in staining solution consisting of 57 ll FluoresceinFragEL TdT Labeling Reaction Mix + 3 ll TdT enzyme solution for 1 h in incubator at 38.5 °C, 5% CO2 in air at maximal humidity. A group of oocytes/ embryos was cultured in 57 ll Fluorescein-FragEL TdT Labeling Reaction Mix without the addition of TdT enzyme solution to serve as control. The oocytes/embryos were mounted onto a microscope slide with a drop of mounting medium containing DABCO (1,4-diazabi-cyclo-2,2,2-octane) as the fluorescent signal-sustaining component and covered with a coverslip. The slide was then observed with a Nikon inverted microscope equipped with fluorescence optics. Photographs were taken with ASA 400 Fuji color film under 200 magnification.
rhodamine fluorophores conjugated on each side of the caspase-3 cleavage site. The fluorescence is quenched in intact substrate due to the folded peptide structure. Upon cleavage by the activity of caspase-3, high intensity red rhodamine fluorescence can be obtained. To perform the assay, oocytes/embryos were cultured in 50 ll RPMI 1640 medium with 10 lM peptide substrate + 10% FBS at 38.5 °C, 5% CO2 in air at maximal humidity. The optimal incubation time (60 min) for both oocytes and embryos was determined by taking samples from the incubation at 30, 60, 90, and 120 min after the onset of incubation and checked under a fluorescence microscope. A portion of degenerating oocytes or embryos was fixed in 4% paraformaldehyde in PBS for 1 h to inactivate caspase-3 prior to its detection treatment. Fixed oocytes or embryos (control 1) and morphologically viable oocytes or embryos (control 2) were also cultured in PhiPhiLux G2 D2 kit for caspase-3 detection. Oocytes or embryos were washed in TL–Hepes for three times to reduce the background fluorescence and were then mounted onto a glass slide with a coverslip. The slides were sealed with clear nail polish and examined under a fluorescence microscope using rhodamine filter combination (EX 510-560, BA 590). Photographs were taken using ASA 400 Fuji color film under 200 magnification. Statistical analysis The experiment was repeated three times. Since oocytes may enter the process of degeneration anytime during culture and the degeneration process is a dynamic process, the results obtained from various assays may not be comparable. Therefore, only the results from viability assay by morphological evaluation and vital stain were compared by v2 test.
Detection of caspase-3 activity Activity of caspase-3 in degenerating oocytes or embryos with fragmented blastomeres was detected using a PhiPhiLux G2 D2 kit (Calbiochem, San Diego, CA). This kit contains a peptide substrate for caspase-3. The substrate GDEVDGI (the caspase cleavage site is underlined) has two
Results Viability assay Immediately after warming, 94% (175/187) of the oocytes were judged by morphological exami-
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nation to have survived the process of vitrification and warming. This result agreed with the result obtained by membrane permeability assay that showed 92% survival rate (172/187; P > 0:05).
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Morphological detection of apoptosis in oocytes and embryos Examination for typical apoptotic morphology was conducted daily after insemination. Cleaved
Fig. 1. Detection of apoptosis in cryopreserved bovine oocytes or embryos derived from cryopreserved oocytes by both morphological characterization, such as condensed cytoplasm (a); Formation of apoptotic bodies (b); and biochemical characterization, such as detection of DNA fragmentation in oocyte (c) and embryo (d); and caspase-3 activity in oocytes (e) and in blastomeres of arrested embryos (f).
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embryos were separated from uncleaved oocytes at 48 h post-insemination. On Day 2, 11% (35/324) of uncleaved oocytes showed cytoplasmic condensation (Fig. 1a). On Day 3, an additional 6% (19/324) of oocytes had condensed cytoplasm. These oocytes were cultured separately. Oocytes with fragmented cytoplasm and apoptotic body formation were not observed until Day 3 of culture. These oocytes accounted for a small proportion (15%, 47/324) of the uncleaved oocytes (Fig. 1b). Other degenerating oocytes either had dark or yellowish cytoplasm without observable cytoplasmic fragmentation (39%, 127/324). The remainder of uncleaved oocytes did not show any sign of degeneration by Day 3. In the meantime, we did not observe any oocytes displaying morphological characteristics of necrosis, such as cell swelling. In embryos derived from cryopreserved oocytes, a high proportion (61%, 174/285) of embryos had arrested development as assessed on Day 4. DNA fragmentation assay The DNA fragmentation assay was conducted on 90 uncleaved oocytes showing signs of degeneration on Day 3. Prior to the assay, degenerating oocytes were separated into three groups: oocytes with dark cytoplasm, fragmented cytoplasm, and yellowish cytoplasm. Positive signals (Fig. 1c) were observed in 79% (33/42) of oocytes with dark cytoplasm and 76% (16/21) of oocytes with fragmented cytoplasm. Oocytes with yellowish cytoplasm were negative for this assay (0/27). Existence of fragmented DNA in blastomere(s) of embryos was examined on Day 4. Fragmented DNA in blastomeres was detected in 78% (54/69) of embryos evaluated (Fig. 1d). In the control group, there were no positive signals for DNA breaks in oocytes (0/26) and embryos (0/22) cultured with only Fluorescein-FragEL TdT Labeling Reaction Mix without the addition of TdT enzyme solution. Caspase-3 activity assay As in the DNA fragmentation assay, 98 Day 3 degenerating oocytes were classified into groups having fragmented, dark or yellowish cytoplasm.
Positive caspase-3 activity was found in 89% (17/ 19), 88% (45/51), and 43% (12/28) of oocytes having fragmented, dark or yellowish cytoplasm, respectively (Fig. 1e). Ninety percent (67/71) of the arrested embryos had blastomeres with caspase-3 activity (Fig. 1f). In the visually assessed viable oocytes and embryos, no positive signal was found (0/25 and 0/13, respectively). No caspase-3 activity was found in paraformaldehyde-fixed degenerating oocytes (0/21) or embryos (0/9).
Discussion Therein, we found strong evidence that cryopreserved oocytes and embryos derived from cryopreserved oocytes degenerate through an apoptotic cell death mechanism after warming and return to culture by combined use of morphological and biochemical assays. A significant feature of oocyte cryopreservation by all current protocols is the compromised developmental competence of oocytes as mostly demonstrated in human and bovine oocytes [3,37]. In this experiment, greater than 90% of oocytes were viable immediately after warming as assessed by morphological evaluation and confirmed by membrane permeability test. Nevertheless, the detrimental effect of cryopreservation was expressed gradually during in vitro culture. The first massive loss of oocytes occurred due to failure in fertilization. Nearly 50% of oocytes lose the ability to cleave [38]. A degeneration mechanism was initiated in those uncleaved oocytes. Among the cleaved embryos, only about 5% could develop into blastocyst stage. The majority stopped development during the first few days of culture and underwent degeneration. Cells normally degenerate via either apoptotic or necrotic pathways. In three replicates of this experiment, we did not observe oocytes having the morphological characteristics of necrosis. Degenerating oocytes did not undergo cell distension, lysis, and leakage of intracellular contents [1,41]; instead, we observed apoptotic morphological changes in some of the cryopreserved oocytes in culture similar to observations in uncryopreserved oocytes reported by others [9,28,29,34], such as
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cytoplasmic condensation, fragmentation of cytoplasm, and the formation of apoptotic bodies. In addition, when assays for biochemical markers of apoptosis, such as DNA fragmentation and caspase-3 activity, were used, we found that the positive signals for these markers were detectable in both oocytes with and without observable apoptotic morphology. In performing these assays, we set-up controls to ensure that the signals we observed were the correct signals for apoptosis. DNA fragmentation was detectable using FluoresceinFragEL in some oocytes with condensed cytoplasm, fragmented cytoplasm, and dark cytoplasm. In embryos, DNA fragmentation was detected in blastomeres of arrested embryos. Also we used a peptide substrate GDEVDGI to detect the caspase-3 activity. We detected caspase-3 activity in high proportions on oocytes from the three groups on Day 3 post-insemination and in fragmented blastomeres in Day 4 embryos. Although we could not eliminate the possibility of necrosis directly by biochemical methods, we provided strong evidence that bovine oocytes and embryos degenerated through apoptosis as their non-cryopreserved counterparts [26,44]. Detection of nuclear DNA breakage by in situ DNA end-labeling technique has been used to detect apoptosis by many laboratories in various cell and tissue types including mammalian oocytes and embryos [1,5,9,10,21,35]. We obtained positive signals for nuclear DNA fragmentation in both oocytes and embryos with a DNA fragmentation detection kit. Moreover, existence of intracellular caspase activity is a definitive confirmation of apoptotic events because caspases are apoptosisspecific proteases. Activated caspases cleave specifically at the carboxyl side of aspartate residues of certain peptide motifs [6]. The only known proteases with this specificity are caspases and cytotoxic T lymphocyte serine protease granzyme B in mammals [7]. In this experiment, we detected caspase-3 activity in degenerating oocytes and embryos at very high proportion. Noticeably, not all oocytes or embryos showing signs of degeneration had characteristics of apoptosis at the time of examination with morphological and biochemical assays. This may be due to the following reasons. First, because of the various degrees of
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cryoinjuries in different oocytes resulting from cryopreservation, apoptosis was not initiated simultaneously in all developmentally impaired oocytes. Instead, oocytes or embryos may enter apoptotic cell death pathway at any time point during culture. Second, we may simply miss the right point for a particular assay in some oocytes or embryos that underwent apoptosis because apoptosis itself is a multistage, active process. Therefore, we believe that the cryopreserved oocytes and embryos derived from cryopreserved oocytes degenerate through apoptosis. The exact mechanisms of cryopreservation in inducing apoptosis need to be investigated. In our experiments, intact oocytes were inseminated after warming. Therefore, the possibility that the initiation of apoptosis in oocytes is due to the lack of continuous development signals, such as the event of fertilization, can be eliminated. Several factors, such as cooling, cryoprotectants, and osmotic insult, may contribute to the initiation of apoptosis. Hypothermia can induce apoptosis in rat germ cells at specific stages and in cultured murine cells [4,17]. The high concentration of cryoprotectants used for vitrification is toxic to cells [37]. Also, cryoprotectants may not be completely removed from cells after warming. The residual cryoprotectants left inside the cells may have the possibility to interact with cellular structures or molecules and result in developmental arrest. Osmotic stresses, both hypo-osmolarity and hyperosmolarity, induce apoptosis in various cell types, such as cardiac fibroblasts [24], fish cells [11], and rat alveolar type II cells [8]. During the process of cryopreservation, these factors could result in various cryoinjuries in oocytes structurally [13,22,32] and/or biochemically and these damages in turn impair the oocyteÕs ability to respond to signals for continued development. As a result, an apoptotic mechanism is initiated to eliminate the developmentally retarded oocytes or embryos. In current practice, we treat cryopreserved oocytes the same way as we treat uncryopreserved oocytes; that is, we employ the same protocols for in vitro maturation, fertilization, and culture. However, the various treatments during cryopreservation may impose some special features on oocytes that cause such oocytes to have less
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Degeneration of cryopreserved bovine oocytes via apoptosis during subsequent cultureq Hongsheng Men,*,1 Rick L. Monson, John J. Parrish, and Jack J. Rutledge Department of Animal Sciences, University of Wisconsin-Madison, 1675 Observatory Drive, Madison, WI 53706, USA Received 21 February 2003; accepted 4 July 2003
Abstract Cryopreservation causes a significant proportion of bovine oocytes to undergo degeneration during subsequent culture. We investigated the degeneration mechanism of cryopreserved oocytes. In vitro matured bovine oocytes were vitrified by the open-pulled straw (OPS) method. In each replicate, a group of oocytes were randomly taken after warming to determine oocyte survival by both morphological evaluation and propidium iodide vital staining. The remainders were evaluated by morphological criterion. Morphologically intact oocytes were co-incubated with frozenthawed spermatozoa for subsequent development. In situ examination of DNA breaks in oocytes and embryos was conducted using a Fluorescein-FragEL DNA fragmentation detection kit. A caspase-3 detection kit was used to detect caspase-3 activity in oocytes and embryos. Most of the oocytes survived cooling and warming processes as assessed by both morphological evaluation and vital stain. During subsequent culture, some degenerating oocytes displayed observable apoptotic morphology, such as cytoplasmic condensation, cytoplasmic fragmentation, and formation of apoptotic bodies. Biochemical markers of apoptosis, such as apoptotic DNA fragmentation and activation of caspases, were detected not only in oocytes having typical apoptotic morphology, but also in oocytes without observable apoptotic morphology. In embryos, positive signals for both biochemical markers were detected in blastomeres. This experiment suggests that cryopreserved bovine oocytes degenerate via apoptosis during subsequent culture. Ó 2003 Elsevier Inc. All rights reserved. Keywords: Apoptotic cell death; Caspase-3 activity; DNA fragmentation; Vitrification
q
This work was supported by the Babcock Institute for International Dairy Research and Development, University of Wisconsin-Madison, and by a grant from USDA/CSREES 2001-52 101-11318. * Corresponding author. Fax: 1-573-8847521. E-mail address: [email protected] (H. Men). 1 Present address: Department of Veterinary Pathobiology, University of Missouri-Columbia, 1600 E. Rollins Street, Columbia, MO 65211, USA.
A dominant feature of mammalian (especially human and domestic animal) oocyte cryopreservation by current protocols is loss of developmental competence. Cryopreservation causes a significant increase in the rate of degeneration in oocytes and embryos derived from cryopreserved oocytes during culture [18,19]. Current efforts in the improvement of oocyte cryopreservation concentrate on increasing the cooling rate to minimize cryoinjuries [20,38]. It may be possible to improve
0011-2240/$ - see front matter Ó 2003 Elsevier Inc. All rights reserved. doi:10.1016/S0011-2240(03)00070-1
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oocyte cryopreservation by inhibiting the initiation of oocyte degeneration during subsequent culture if the degeneration mechanism of cryopreserved oocytes can be characterized. Cells normally degenerate by one of two mechanisms: apoptosis and necrosis. Apoptosis is an endogenous cell degeneration mechanism that is necessary for normal development and tissue homeostasis [43]. The apoptotic process can also be activated in response to a wide variety of nonphysiological stimuli, such as temperature, toxicants, oxidative stress, and X-radiation [41]. Morphological characteristics of apoptotic cells are cell blebbing, margination of the chromatin, and the formation of apoptotic bodies (membranebounded cytoplasmic bodies) [42]. These morphological changes are a result of the activation of a group of proteases, such as caspase-1, caspase-2, caspase-3, etc. Caspases exist in cells as inactive proenzymes. Upon apoptosis initiation, they are activated by cleavage at specific aspartate residues [6]. A variety of proteins, such as PARP, histone H1, DNA-dependent protein kinase, and lamins are destroyed by the action of caspases and result in the morphological changes in apoptotic cells. Generation of DNA fragments of 180–200 bp, a characteristic of apoptosis, is a result of caspaseactivated endonucleases [23,42,43]. On the other hand, when cells experience a wide departure from physiological conditions, such as injury or trauma, another mechanism, necrosis, takes over as the mechanism for cell degeneration. Necrotic cell death is characterized by cell distension followed by lysis and leakage of intracellular contents. Nuclear DNA is randomly cleaved and forms a ‘‘smear’’ in agarose gel after electrophoresis [1,41]. During germ cell development, apoptosis also plays a significant role in germ cell homeostasis. Apoptotic cell death can be observed as early as the primordial germ cell proliferation stage [40] and continues throughout spermatogenesis [15,30] and oogenesis [31,36]. Up to 75% of male germ cells are lost during spermatogenesis [30]. During ovarian development, over two-thirds of oocytes are eliminated through apoptosis before birth and a large number of remaining oocytes are lost through apoptosis during follicular atresia [36].
Ovulated oocytes also undergo degeneration if fertilization does not occur. It is generally thought that apoptosis is the degeneration mechanism of ovulated oocytes [9,28,34] although Van Blerkom and Davis [39] present evidence otherwise. In vitro cultured mammalian (including bovine) oocytes and embryos without cryopreservation are also degenerated through an apoptotic pathway [10,12,14,26,44]. However, the investigation into the mechanism behind degeneration of cryopreserved oocytes is lacking. It is essential to elucidate the mechanism behind oocyte degeneration after cryopreservation, because cryopreserved oocytes are currently fertilized and cultured to the blastocyst stage in vitro using the same culture procedures developed for non-cryopreserved oocytes. However, there are reasons that cryopreserved oocytes may need a specialized culture system. During cryopreservation, oocytes experience severe adverse physiological conditions. These adverse conditions render oocytes with various cytological injuries [3,32]. In addition to cytological abnormalities, cryopreservation may also result in biochemical abnormalities to oocytes as it has to other cell types. For example, in bone marrow cells, cryopreservation can activate intracellular proteases and the activated proteases in turn cause artificial cleavage of apoptosis-related proteins [33]. Therefore, prevention of the initiation of apoptosis would in turn increase survival of cryopreserved cells. Many cell types are known to require modification of freezing solutions or culture systems for optimal growth or survival. Apoptotic inhibitors in the cryopreservation solution or cold storage solution have enhanced the survival of several cell lines or tissue [2,16,33]. Vascular endothelial growth factor shows a protective effect on cold preserved sinusoidal endothelial cells during storage and culture [25]. Therefore, it would be advantageous in guiding the cryopreservation or the design of culture system for cryopreserved oocytes if we can characterize the degeneration mechanism of cryopreserved oocytes and embryos derived from cryopreserved oocytes. Herein, we test the hypothesis that cryopreserved oocytes and embryos derived from cryopreserved oocytes degenerate via apoptosis.
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Materials and methods Unless otherwise indicated, all chemicals were purchased from Sigma Chemical (St. Louis, MO). Maturation, fertilization, and culture of bovine oocytes In this study, we employed a bovine oocyte maturation, fertilization, and culture system that could consistently give 80% cleavage rate and 30% blastocyst formation rate (out of cleaved embryos). Cumulus–oocyte complexes were obtained by aspiration of 3–8 mm follicles from ovaries collected from a local slaughterhouse. Oocytes with uniformly granulated cytoplasm and at least one layer of cumulus cell investment were washed three times in TL–Hepes (BioWhittaker, Walkersville, MD) and matured for 22 h at 38.5 °C, 5% CO2 in air with maximal humidity in 50 ll drops of medium TCM-199 (BioWhittaker) with the following supplementations: 10% FBS (Gibco-BRL, Grand Island, NY), 0.2 mM Na-pyruvate, 3 lg/ml FSH (National Institutes of Health, Bethesda, MD), 3 lg/ml LH (National Institutes of Health), 1 lg/ml estradiol, and 25 lg/ml gentamycin. Oocytes were fertilized with 45–90% Percoll gradient selected living sperm from frozen semen at a final concentration of 1 106 sperm/ml in a 50 ll fertilization medium drop under 5% CO2 in air with maximal humidity at 38.5 °C [27]. The fertilization medium used is a modified TyrodeÕs medium supplemented with 0.2 mM Na-pyruvate, 6 mg/ml fatty acid free BSA, 2 lg/ml heparin, 20 lM penicillamine, 10 lM hypotaurine, and 1 lM/ml epinephrine. After 22 h of fertilization, putative zygotes were cultured in 50 ll drops of KSOM (Specialty Media, MD) with 3 mg/ml BSA as the protein source. Vitrification of bovine oocytes Protocols of oocyte vitrification with openpulled straws (OPS) were as described by Vajta et al. [38]. After partial removal of cumulus cells, mature oocytes were first dehydrated by 45 s exposure to 10% (v/v) ethylene glycol (EG) + 10% (v/v) dimethyl sulfoxide (Me2 SO) + TL–Hepes with
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20% (v/v) FBS supplementation and then in vitrification solution (20% EG + 20% Me2 SO + 0.5 M sucrose in TL–Hepes + 20% FBS). Oocytes with minimal amount (1–2 ll) of vitrification solution were picked up in the narrow end of the OPS by capillary action. The straw was plunged directly into liquid nitrogen within 30 s of contacting the concentrated vitrification solution. Vitrified oocytes were stored in liquid nitrogen for 1 or 2 days. Warming was conducted by immersing the narrow end of the OPS into TL–Hepes with 0.25 M sucrose and 20% FBS for 1 min at 37 °C. Rehydration was conducted by subsequent washes for 5 min in each wash: once in TL–Hepes with 0.15 M sucrose + 20% FBS and then twice in TL–Hepes + 20% FBS. Viability assay Viability of warmed oocytes was determined based on visual examination of the integrity of the oocyte membrane, zona pellucida, and the normality of the cytoplasm immediately after warming. The validity of morphological classification was confirmed by vital staining with propidium iodide. To perform vital stain, samples of visually viable and dead oocytes were mounted separately on microscope slides and then covered by coverslips with the support of Vaseline–wax mix. Oocytes were stained with propidium iodide at a final concentration of 0.2 mg/ml in PBS. The oocytes were examined under a Nikon Diaphot microscope equipped with epifluorescent microscopy. A combination of 510–560 nm excitation filter and 590 nm barrier filter was used to examine PI fluorescence. The remaining oocytes were evaluated by morphological observation. Visually dead oocytes were discarded, and the morphologically intact oocytes were subject to fertilization and culture. Morphological characterization of apoptosis in oocytes Morphological characterization of apoptosis was carried out under a Bausch and Lomb stereomicroscope. Oocytes having the observable features of apoptotic morphology, such as condensed cytoplasm, fragmented cytoplasm and
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apoptotic bodies were considered to undergo apoptotic cell death. Photographs were taken with a Zeiss inverted microscope equipped with differential interference contrast optics using ASA 400 Fuji black and white film with 200 magnification. Detection of DNA fragmentation in bovine oocytes and embryos DNA integrity of degenerating oocytes or embryos with fragmented blastomeres was evaluated by using DNA fragmentation detection kit (Fluorescein-FragEL, Oncogene Research Products, Boston, MA). The principle of Fluorescein-FragEL is that terminal deoxynucleotidyl transferase (TdT) catalyzes the addition of fluorescein-labeled and unlabeled deoxynucleotides to the 30 -OH ends generated by endonucleases during apoptotic degeneration. Oocytes or embryos were first fixed in 4% paraformaldehyde in PBS + 1% BSA for 15 min and permeabilized for 5 min in 0.01% Triton X-100 in PBS–BSA at room temperature. After three washes in PBS–BSA, oocytes/embryos were incubated in staining solution consisting of 57 ll FluoresceinFragEL TdT Labeling Reaction Mix + 3 ll TdT enzyme solution for 1 h in incubator at 38.5 °C, 5% CO2 in air at maximal humidity. A group of oocytes/ embryos was cultured in 57 ll Fluorescein-FragEL TdT Labeling Reaction Mix without the addition of TdT enzyme solution to serve as control. The oocytes/embryos were mounted onto a microscope slide with a drop of mounting medium containing DABCO (1,4-diazabi-cyclo-2,2,2-octane) as the fluorescent signal-sustaining component and covered with a coverslip. The slide was then observed with a Nikon inverted microscope equipped with fluorescence optics. Photographs were taken with ASA 400 Fuji color film under 200 magnification.
rhodamine fluorophores conjugated on each side of the caspase-3 cleavage site. The fluorescence is quenched in intact substrate due to the folded peptide structure. Upon cleavage by the activity of caspase-3, high intensity red rhodamine fluorescence can be obtained. To perform the assay, oocytes/embryos were cultured in 50 ll RPMI 1640 medium with 10 lM peptide substrate + 10% FBS at 38.5 °C, 5% CO2 in air at maximal humidity. The optimal incubation time (60 min) for both oocytes and embryos was determined by taking samples from the incubation at 30, 60, 90, and 120 min after the onset of incubation and checked under a fluorescence microscope. A portion of degenerating oocytes or embryos was fixed in 4% paraformaldehyde in PBS for 1 h to inactivate caspase-3 prior to its detection treatment. Fixed oocytes or embryos (control 1) and morphologically viable oocytes or embryos (control 2) were also cultured in PhiPhiLux G2 D2 kit for caspase-3 detection. Oocytes or embryos were washed in TL–Hepes for three times to reduce the background fluorescence and were then mounted onto a glass slide with a coverslip. The slides were sealed with clear nail polish and examined under a fluorescence microscope using rhodamine filter combination (EX 510-560, BA 590). Photographs were taken using ASA 400 Fuji color film under 200 magnification. Statistical analysis The experiment was repeated three times. Since oocytes may enter the process of degeneration anytime during culture and the degeneration process is a dynamic process, the results obtained from various assays may not be comparable. Therefore, only the results from viability assay by morphological evaluation and vital stain were compared by v2 test.
Detection of caspase-3 activity Activity of caspase-3 in degenerating oocytes or embryos with fragmented blastomeres was detected using a PhiPhiLux G2 D2 kit (Calbiochem, San Diego, CA). This kit contains a peptide substrate for caspase-3. The substrate GDEVDGI (the caspase cleavage site is underlined) has two
Results Viability assay Immediately after warming, 94% (175/187) of the oocytes were judged by morphological exami-
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nation to have survived the process of vitrification and warming. This result agreed with the result obtained by membrane permeability assay that showed 92% survival rate (172/187; P > 0:05).
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Morphological detection of apoptosis in oocytes and embryos Examination for typical apoptotic morphology was conducted daily after insemination. Cleaved
Fig. 1. Detection of apoptosis in cryopreserved bovine oocytes or embryos derived from cryopreserved oocytes by both morphological characterization, such as condensed cytoplasm (a); Formation of apoptotic bodies (b); and biochemical characterization, such as detection of DNA fragmentation in oocyte (c) and embryo (d); and caspase-3 activity in oocytes (e) and in blastomeres of arrested embryos (f).
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embryos were separated from uncleaved oocytes at 48 h post-insemination. On Day 2, 11% (35/324) of uncleaved oocytes showed cytoplasmic condensation (Fig. 1a). On Day 3, an additional 6% (19/324) of oocytes had condensed cytoplasm. These oocytes were cultured separately. Oocytes with fragmented cytoplasm and apoptotic body formation were not observed until Day 3 of culture. These oocytes accounted for a small proportion (15%, 47/324) of the uncleaved oocytes (Fig. 1b). Other degenerating oocytes either had dark or yellowish cytoplasm without observable cytoplasmic fragmentation (39%, 127/324). The remainder of uncleaved oocytes did not show any sign of degeneration by Day 3. In the meantime, we did not observe any oocytes displaying morphological characteristics of necrosis, such as cell swelling. In embryos derived from cryopreserved oocytes, a high proportion (61%, 174/285) of embryos had arrested development as assessed on Day 4. DNA fragmentation assay The DNA fragmentation assay was conducted on 90 uncleaved oocytes showing signs of degeneration on Day 3. Prior to the assay, degenerating oocytes were separated into three groups: oocytes with dark cytoplasm, fragmented cytoplasm, and yellowish cytoplasm. Positive signals (Fig. 1c) were observed in 79% (33/42) of oocytes with dark cytoplasm and 76% (16/21) of oocytes with fragmented cytoplasm. Oocytes with yellowish cytoplasm were negative for this assay (0/27). Existence of fragmented DNA in blastomere(s) of embryos was examined on Day 4. Fragmented DNA in blastomeres was detected in 78% (54/69) of embryos evaluated (Fig. 1d). In the control group, there were no positive signals for DNA breaks in oocytes (0/26) and embryos (0/22) cultured with only Fluorescein-FragEL TdT Labeling Reaction Mix without the addition of TdT enzyme solution. Caspase-3 activity assay As in the DNA fragmentation assay, 98 Day 3 degenerating oocytes were classified into groups having fragmented, dark or yellowish cytoplasm.
Positive caspase-3 activity was found in 89% (17/ 19), 88% (45/51), and 43% (12/28) of oocytes having fragmented, dark or yellowish cytoplasm, respectively (Fig. 1e). Ninety percent (67/71) of the arrested embryos had blastomeres with caspase-3 activity (Fig. 1f). In the visually assessed viable oocytes and embryos, no positive signal was found (0/25 and 0/13, respectively). No caspase-3 activity was found in paraformaldehyde-fixed degenerating oocytes (0/21) or embryos (0/9).
Discussion Therein, we found strong evidence that cryopreserved oocytes and embryos derived from cryopreserved oocytes degenerate through an apoptotic cell death mechanism after warming and return to culture by combined use of morphological and biochemical assays. A significant feature of oocyte cryopreservation by all current protocols is the compromised developmental competence of oocytes as mostly demonstrated in human and bovine oocytes [3,37]. In this experiment, greater than 90% of oocytes were viable immediately after warming as assessed by morphological evaluation and confirmed by membrane permeability test. Nevertheless, the detrimental effect of cryopreservation was expressed gradually during in vitro culture. The first massive loss of oocytes occurred due to failure in fertilization. Nearly 50% of oocytes lose the ability to cleave [38]. A degeneration mechanism was initiated in those uncleaved oocytes. Among the cleaved embryos, only about 5% could develop into blastocyst stage. The majority stopped development during the first few days of culture and underwent degeneration. Cells normally degenerate via either apoptotic or necrotic pathways. In three replicates of this experiment, we did not observe oocytes having the morphological characteristics of necrosis. Degenerating oocytes did not undergo cell distension, lysis, and leakage of intracellular contents [1,41]; instead, we observed apoptotic morphological changes in some of the cryopreserved oocytes in culture similar to observations in uncryopreserved oocytes reported by others [9,28,29,34], such as
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cytoplasmic condensation, fragmentation of cytoplasm, and the formation of apoptotic bodies. In addition, when assays for biochemical markers of apoptosis, such as DNA fragmentation and caspase-3 activity, were used, we found that the positive signals for these markers were detectable in both oocytes with and without observable apoptotic morphology. In performing these assays, we set-up controls to ensure that the signals we observed were the correct signals for apoptosis. DNA fragmentation was detectable using FluoresceinFragEL in some oocytes with condensed cytoplasm, fragmented cytoplasm, and dark cytoplasm. In embryos, DNA fragmentation was detected in blastomeres of arrested embryos. Also we used a peptide substrate GDEVDGI to detect the caspase-3 activity. We detected caspase-3 activity in high proportions on oocytes from the three groups on Day 3 post-insemination and in fragmented blastomeres in Day 4 embryos. Although we could not eliminate the possibility of necrosis directly by biochemical methods, we provided strong evidence that bovine oocytes and embryos degenerated through apoptosis as their non-cryopreserved counterparts [26,44]. Detection of nuclear DNA breakage by in situ DNA end-labeling technique has been used to detect apoptosis by many laboratories in various cell and tissue types including mammalian oocytes and embryos [1,5,9,10,21,35]. We obtained positive signals for nuclear DNA fragmentation in both oocytes and embryos with a DNA fragmentation detection kit. Moreover, existence of intracellular caspase activity is a definitive confirmation of apoptotic events because caspases are apoptosisspecific proteases. Activated caspases cleave specifically at the carboxyl side of aspartate residues of certain peptide motifs [6]. The only known proteases with this specificity are caspases and cytotoxic T lymphocyte serine protease granzyme B in mammals [7]. In this experiment, we detected caspase-3 activity in degenerating oocytes and embryos at very high proportion. Noticeably, not all oocytes or embryos showing signs of degeneration had characteristics of apoptosis at the time of examination with morphological and biochemical assays. This may be due to the following reasons. First, because of the various degrees of
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cryoinjuries in different oocytes resulting from cryopreservation, apoptosis was not initiated simultaneously in all developmentally impaired oocytes. Instead, oocytes or embryos may enter apoptotic cell death pathway at any time point during culture. Second, we may simply miss the right point for a particular assay in some oocytes or embryos that underwent apoptosis because apoptosis itself is a multistage, active process. Therefore, we believe that the cryopreserved oocytes and embryos derived from cryopreserved oocytes degenerate through apoptosis. The exact mechanisms of cryopreservation in inducing apoptosis need to be investigated. In our experiments, intact oocytes were inseminated after warming. Therefore, the possibility that the initiation of apoptosis in oocytes is due to the lack of continuous development signals, such as the event of fertilization, can be eliminated. Several factors, such as cooling, cryoprotectants, and osmotic insult, may contribute to the initiation of apoptosis. Hypothermia can induce apoptosis in rat germ cells at specific stages and in cultured murine cells [4,17]. The high concentration of cryoprotectants used for vitrification is toxic to cells [37]. Also, cryoprotectants may not be completely removed from cells after warming. The residual cryoprotectants left inside the cells may have the possibility to interact with cellular structures or molecules and result in developmental arrest. Osmotic stresses, both hypo-osmolarity and hyperosmolarity, induce apoptosis in various cell types, such as cardiac fibroblasts [24], fish cells [11], and rat alveolar type II cells [8]. During the process of cryopreservation, these factors could result in various cryoinjuries in oocytes structurally [13,22,32] and/or biochemically and these damages in turn impair the oocyteÕs ability to respond to signals for continued development. As a result, an apoptotic mechanism is initiated to eliminate the developmentally retarded oocytes or embryos. In current practice, we treat cryopreserved oocytes the same way as we treat uncryopreserved oocytes; that is, we employ the same protocols for in vitro maturation, fertilization, and culture. However, the various treatments during cryopreservation may impose some special features on oocytes that cause such oocytes to have less
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tolerance to current in vitro culture systems. Special treatment, such as inhibition of proteases, may be needed for culture of cryopreserved oocytes. Inclusion of an inhibitor (caspase I inhibitor V) in the cryopreservation solution was effective in reducing post-cryopreservation apoptotic degeneration of a canine renal cell line [2]. Growth factors may also have a protective effect on cooling-induced apoptotic cell degeneration, such as vascular endothelial growth factor on cold preserved sinusoidal endothelial cells during storage and culture [25]. In this experiment, we provided strong evidence that the degeneration mechanism of cryopreserved oocytes is apoptosis. Therefore, inhibition of apoptosis during cryopreservation and subsequent culture, or stimulation of the oocyte mitogenic potential could be approaches of choice in the improvement of mammalian oocyte cryopreservation. References [1] R.T Allen, W.J. Hunter III, D.K. Agrawal, Morphological and biochemical characterization and analysis of apoptosis, J. Pharmacol. Toxicol. Methods 37 (1997) 215–228. [2] J.M. Baust, R. Van Buskirk, J.G. Baust, Cell viability improves following inhibition of cryopreservation-induced apoptosis, In Vitro Cell Dev. Biol. Animal 36 (2000) 262–270. [3] B. Bernard, B.J. Fuller, Cryopreservation of human oocytes: a review of current problems and perspectives, Hum. Reprod. Update 2 (1996) 193–207. [4] J. Blanco-Rodriguez, C. Martinez-Garcia, Mild hypothermia induces apoptosis in rat testis at specific stages of the seminiferous epithelium, J. Androl. 18 (1997) 535–539. [5] A.T. Byrne, J. Southgate, D.R. Brison, H.J. Leese, Analysis of apoptosis in the preimplantation bovine embryo using TUNEL, J. Reprod. Fert. 117 (1999) 97–105. [6] G.M. Cohen, Caspases: the executioners of apoptosis, Biochem. J. 326 (1997) 1–16. [7] W. Earnshaw, L.M. Martins, S.H. Kaufmann, Mammalian caspases: structure, activation, substrates, and functions during apoptosis, Annu. Rev. Biochem. 68 (1999) 383–424. [8] Y.S. Edwards, L.M. Sutherland, J.H.T. Power, T.E. Nicholas, A.W. Murray, Osmotic stress induces both secretion and apoptosis in rat alveolar type II cells, Am. J. Physiol. 275 (1998) L670–L678. [9] Y. Fujino, K. Ozaki, S. Yamamasu, F. Ito, I. Matsuoka, E. Hayashi, H. Nakamura, S. Ogita, E. Sato, M. Inoue,
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