- Email: [email protected]
Gamete origin in relation to early embryo development
Theriogenology 68S (2007) S131–S137 www.theriojournal.com
Gamete origin in relation to early embryo development A. Van Soom a,*, L. Vandaele a, K. Goossens b, A. de Kruif a, L. Peelman b a
Reproductive Biology Unit, Department of Reproduction, Obstetrics and Herd Health, Faculty of Veterinary Medicine, Ghent University, Salisburylaan 133, B-9820 Merelbeke, Belgium b Molecular Biology Unit, Heidestraat 19, Faculty of Veterinary Medicine, 9820 Merelbeke, Belgium
Abstract Fertilization in vivo requires a complex series of selection events to occur in order to guarantee that only the fittest gametes take part in the fusion process and give rise to a viable embryo. Conventional practice in bovine in vitro fertilization however is to select oocytes and sperm by quite crude procedures. It is therefore not inconceivable that essentially unfit gametes may drive aberrant embryo development in vitro. Abnormal embryonic cells are being removed by apoptosis, which is a physiological process in embryos. Only an excess or a lack of apoptosis can lead to embryonic death or abnormal development. Suboptimal culture conditions undoubtedly contribute to undue embryonic apoptosis, but the intrinsic quality of the oocyte may also be a causative factor. It is generally accepted that the oocyte is in control of early embryogenesis, but is it also in control of future embryonic suicide? Is a compromised follicular environment predestining the oocyte to a dire fate? What is the contribution of the cumulus cells to oocyte quality, and can they rescue it from early demise? And what can be said about the origin of the spermatozoa? Research in human in vitro fertilization has definitely shown that factors such as paternal age, smoking and other sperm stressors can contribute to abnormal embryo development and even diseased offspring. This review will address the questions raised above, and will describe what is known about the cellular and molecular biology that may account for abnormal bovine embryo development caused by gamete origin. # 2007 Elsevier Inc. All rights reserved. Keywords: Gamete origin; Embryo; Apoptosis
1. Introduction A tantalizing phrase which has been introduced for the first time in Darwin’s Origin of species is ‘‘survival of the fittest’’. Since the title of this talk reminded me in some way to evolutionary biology, I wondered whether this very concept could not be used to explain embryonic development as well. In modern biology, the term fitness measures reproductive success and is not explicit about the specific ways in which organisms (or in our case, gametes or cells) can be ‘‘fit’’ as in
* Corresponding author. Tel.: +32 9 264 7561; fax: +32 9 264 7797. E-mail address: [email protected] (A. Van Soom). 0093-691X/$ – see front matter # 2007 Elsevier Inc. All rights reserved. doi:10.1016/j.theriogenology.2007.03.019
‘‘having phenotypic characteristics which enhance survival and reproduction’’. The methods we use to select the gametes meant for assisted reproduction and the subsequent treatments we inflict on them are surely imposing a kind of selection pressure, albeit not a natural but merely an artificial one. Natural selection is the central concept of Darwinian theory – the fittest survive, and can give rise to offspring – but do we actually select the fittest gametes? Which are the fittest? And how is ‘‘fitness’’ defined? The study of links between development and evolution has been relatively neglected by animal scientists and veterinarians, who prefer to focus their scientific concepts on the animal as a whole, or to find potential applications for a certain technique [1]. The applied embryology that we are
S132
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137
practicing does not aim to analyze evolutionary mechanisms, since there is no reason to believe that these mechanisms are involved in the process of embryo development in vitro. But can we be sure that this is really so? Are not there numerous examples in assisted reproductive biology to prove that deliberate selection of gametes and/or embryos is causing an effect on the actual outcome? We probably all agree that embryo development is being affected by gamete origin. Gamete origin can be defined at different levels: it can relate to the donor, to the in vivo or in vitro maturation status of the gamete, and to its supporting environment. When these determinants of gamete origin are inferior, they will predestine the resulting embryo for an ill-fated future. Such future can be on short-term aberrant fertilization and abnormal embryo development, resulting in developmental block, retarded development or embryonic demise, or, on the long term, lack of implantation and diseased offspring [2,3]. In this review, we will focus on how we currently define which gametes are fit to take part in the process of reproduction, what the impact of their origin may have on subsequent early embryonic development, and what role apoptosis or programmed cell death plays in determining the fate of the embryo. We hypothesize that the basic machinery to cause embryonic apoptosis may already be present in the oocyte or the sperm. We will review data from the literature and present experimental evidence, obtained in our laboratory mainly in cattle, to support this hypothesis. 2. Of follicles, oocytes and apoptosis Many more follicles are present in the mammalian ovary than will eventually ovulate [4]. Solid evidence is available that the massive loss of germ cells during oogenesis is caused by oocyte apoptosis. Therefore, the normal fate for female germ cells during oogenesis is death. Interestingly, once an antrum has been formed, follicular atresia is mainly driven by apoptosis of granulosa cells, and the oocyte itself is quite resistant to cell death [4]. In fact it can even protect the surrounding cumulus cells against apoptosis [5]. This does not mean, however, that the oocyte is not capable of initiating apoptosis. Transcripts of caspases-3, -7, -8 and -9 have been demonstrated in bovine [6] and mouse oocytes [7], showing that unfertilized oocytes have the machinery to undergo apoptosis by using either the extrinsic (caspase-8 dependent) or intrinsic (caspase-9 dependent) pathways. The different balance in expression of pro- and anti-apoptotic genes may shift the oocyte’s
developmental potential towards either cell death or cell survival, and may be a crucial factor in determining the quality of the oocyte. But how can we predict which oocyte will give rise to a viable embryo and which to a failing one, and can we influence the fitness of the oocyte by means of better maturation conditions? Several aspects of oocyte origin can influence eventual fertilization outcome. During follicular development, oocytes and follicles are forming functional units. This is especially true if we look at the cumulus–oocyte interactions during the late stages of ovogenesis, including oocyte capacitation and maturation. Whereas the intrinsic quality of an oocyte is determined by the presence of a normal set of chromosomes and by its mitochondrial genome, the microenvironment of the oocyte, be it either the preovulatory follicle or the in vitro maturation conditions can influence, on the other hand, the fate of the transcripts and proteins accumulated in the oocyte [8–10]. 2.1. Oocytes unfit to sustain embryo development Are we at risk to select unfit oocytes, when we do apply the morphological criteria which most scientists use very rigorously? The answer is yes. It has long been accepted that the population of oocytes which is being used for in vitro embryo production is a very heterogeneous one. Immature oocytes are being aspirated from antral follicles which are typically between 2 and 8 mm in diameter [11]. A lower developmental potential has already been demonstrated for oocytes derived from smaller follicles (<4 mm) versus those from larger follicles (>6 mm) although no differences could be detected directly at the oocyte level, as far as global profile of transcripts, the pattern of protein neosynthesis and the kinetics of meiosis resumption was concerned [12]. However, it has been demonstrated that bovine follicles smaller than 3 mm contain oocytes, which are smaller than 110 mm [13]. Reaching a critical diameter of 110 mm coincides, for the bovine oocyte at least, with the acquisition of full competence for completing maturation to metaphase II [14]. Embryos generated by smaller oocytes (<110 mm) displayed a higher incidence of apoptosis throughout embryo development as assessed by fluorescent detection of active caspase-3 and -7 in cleaved embryos and morulae [15]. This effect was present until 117 hpi, and may indicate that such embryos are derived from oocytes, which do not possess the complete protein synthetic machinery to prevent embryonic apoptosis. About one third of bovine oocytes possess the transcripts for caspases, but active caspases
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137
are not detectable yet [6]. If however inactive caspases are activated in unfit oocytes, fertilization of such oocytes may lead to embryo fragmentation and impaired embryonic development, because the oocyte cytoplasm cannot provide the necessary transcripts and proteins for adequate suppression of apoptotic pathways [16,17]. Evidence to support this is provided by the finding that oocytes smaller than 110 mm showed in 80% of the cases DNA fragmentation before maturation, whereas this was only 20% for oocytes of larger diameters [18]. More evidence for the importance of follicular origin in determining embryonic fate, is the finding that larger antral follicles are required (>6–8 mm) for collection of oocytes which have the competence for both nuclear and cytoplasmic maturation [19]. Such follicles can be either atretic or healthy in cattle: many more apoptotic granulosa cells are present in the atretic follicles than in the healthy ones, whilst no apoptotic cells could be demonstrated in the cumulus–oocyte complexes from the same follicles [20]. Matters are complicated by the fact that oocytes from early atretic follicles display increased developmental competence [21], and oocyte removal from the follicular environment may artificially induce cumulus cell apoptosis, since cumulus–oocyte complexes show a spontaneous onset of apoptosis during in vitro maturation [20,22], or during prolonged manipulation, predominantly in the outer layers of the cumulus cells (Anguita, unpublished results). How important are these apoptotic cumulus cells for the fate of the oocyte? It has been argued that a high prevalence of apoptotic cumulus cells surrounding the oocyte may be predictive for low developmental potential, in human [23] and in cattle studies [24]. Cumulus cells communicate with each other and with the oocyte via gap junctions (reviewed by [25]), which are essential for transport of nutrients and regulatory factors, but these contacts are lost during the process of maturation. Anyhow, components of follicular fluid, which have been shown to be detrimental for oocyte developmental capacity, such as palmitic and stearic acid, caused increased levels of apoptosis in cumulus cells [10], which is indicative for a causal relationship. On the other hand, growth factors, such as midkine, which have been shown to be beneficial to oocyte developmental competence, have an anti-apoptotic effect on cumulus cells and can exert their effect even through isolated cumulus cells [26]. 2.2. Oocytes fit to sustain embryo development A second maybe much more important question we must ask ourselves is which oocytes are actually fit for
S133
normal embryo production. This question is far more difficult to answer, because no reliable non-invasive methods for oocyte selection are at present available, besides the detection of glucose-6-phosphate dehydrogenase by means of brilliant cresyl blue staining, as recently been reviewed by Ref. [9]. They also stated that analysis of factors in follicular fluid/cells may hold promise to foretell oocyte quality on an individual basis. Probably the most promising approaches in this field are the use of functional genomics, transcriptomics and proteomics, since fit oocytes need to contain a sufficient stockpile of mRNA and proteins to allow embryo development to occur until after embryonic genome activation. First of all, fully-grown oocytes from larger follicles are definitely better equipped to generate embryos. The higher embryo yield of oocytes from larger follicles has been related directly to differences in mRNA transcripts in a recent study [27]. By using the powerful approach of suppressive subtractive hybridization, it was demonstrated that several genes are differentially expressed according to different follicular sizes. Genes involved in cell cycle regulation such as cyclin dependent kinase subunit 1 (CKS1B), pituitary tumor-transforming 1 (PTTG1) and cyclin B2 (CCNB2), and histone 2A (H2A) which contributes to the chromatin support in the early embryo, are gradually upgraded according to follicular size [27]. Second, it would be quite logical to assume that preovulatory follicular fluid provides the best environment for the maturing oocyte. For bovine oocytes, undiluted follicular fluid is not the optimal maturation environment [28], whereas it is added as a routine during pig oocyte maturation. We have recently conducted a number of experiments to get a better understanding of the role of follicular fluid on pig oocyte maturation and found that components larger than 10 kDa have a positive effect on nuclear maturation, but were not able to show an effect on cytoplasmic maturation [29]. Preovulatory porcine follicular fluid however was superior over autologous serum to sustain pig cytoplasmic maturation (Bijttebier, unpublished results). Further studies using proteomic analysis are in progress to identify the molecules involved. Recent data in cattle have also shown that low IGFBP-levels in follicular fluid can be used to select fit oocytes [30]. It remains to be determined whether addition of these molecules to maturation medium can rescue oocytes from early demise. A third determinant of oocyte competence is early cleavage [31,32]. One approach is to use transcriptomics to differentiate between fast and slow cleavers,
S134
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137
and again higher levels of histone H2A [33] and histone H3A [32] have been detected in embryos derived from fit oocytes, which is logical because histones are needed to replace the protamines of the sperm nucleus after fertilization and to assemble embryonic DNA into chromatin for the first cleavages. Other transcripts which were differentially expressed in fast cleavers were stem-loop binding protein (SLBP), which is involved in histone stabilisation [34], YEAF, which is a transcriptional repressor, and isocitrate dehydrogenase (IDH), which modulates oxidative damage [33]. Another approach, which we have used repeatedly in the past, was to look at quality of embryos derived from either fit (fast cleavers) or unfit (slow) oocytes. We found that fast cleavers had higher chances to become a morula or blastocyst, and to display an excellent morphology [31,35]. Moreover, fast cleavers had significantly lower apoptotic indices at 45, 80 and 117 hpi, indicating a maternal effect [15]. The lower incidence of apoptosis may be attributed to the superior stockpile of mRNAs and proteins, which typecast such fit oocytes, but may also be influenced by mitochondria present in fit oocytes. It is generally accepted that the oocyte is in control of early embryogenesis, and our data confirm that it is at least partially in control of future embryonic suicide. Selection of fit oocytes must be followed by optimal insemination and culture conditions to prevent apoptosis to occur. 3. Of spermatozoa, DNA damage and fertilization It has been said that the engine of evolution is the testis [36], with its continuous production of large numbers of spermatozoa by a spermatogenic process that has no means of selecting against phenotypic or genotypic abnormalities. Nature, however, prevents the procreation of severe mutations in the male germ line by selecting against structurally deviant spermatozoa, which have a lower chance to reach and fertilize a mature oocyte [37]. In addition, the sperm mitochondria, which may also carry mutations, are destroyed in the oocyte after fertilization [38]. Two types of DNA damage are however typical for the male germ line: replication errors and DNA fragmentation [39]. Replication errors are more prevalent in the spermatozoa of older men and have been linked with the occurrence of dominant genetic disease, but may be less important for domestic animals, whose lifespan is shorter. DNA damage on the contrary, may occur frequently in ejaculates of otherwise healthy animals, and may arise from three sources: (1) oxidative
stress, (2) abortive Fas-mediated apoptosis and (3) deficiencies in chromatin packaging leading to DNA strand breaks [39]. Fortunately, DNA repair mechanisms which rely on the maternal mRNAs and proteins stored in the oocyte are able to mend errors in DNA, leading to healthy offspring [40]. If an oocyte is fertilized with DNA-damaged spermatozoa, G1-associated DNA repair mechanisms become activated, leading to a dramatic suppression of pronuclear DNA-synthesis via a p53dependent mechanism. Mice lacking such DNA repair enzymes are not viable with preimplantation embryonic death as a result [40]. Which characteristics are needed for spermatozoa to sustain embryo development? It is has been demonstrated that bulls can differ in embryo generating capacity, even if they display similar fertilization rates in vitro. In the early years of human in vitro fertilization, it was postulated that sperm quality and early embryonic development were independent, but later on this was reviewed in favour of a paternal effect on early embryonic viability and further development [41]. Now there is a consensus that suboptimal spermatozoa have a negative influence on the integrity and the quality of the paternal DNA, and may lead to defective embryonic development or diseased offspring. For instance, failure of Sertoli cells to remove sufficient residual cytoplasm from the spermatozoa during spermatogenesis enhances free radical production in the ejaculates of infertile human patients [39]. Evidence exists that children from heavy smokers, whose semen is characterized by oxidative DNA base damage, are four to five times more likely to develop childhood cancer than children from non-smokers [42]. 3.1. Spermatozoa unfit to sustain embryo development Differences in development between embryos sired by different bulls can first be detected before cleavage to the two-cell stage (but after pronuclear formation) and thus before expression of the embryonic genome. Zygotes sired by low fertility bulls showed a delay in Sphase initiation compared to zygotes sired by highfertility bulls [43]: it has been hypothesized that sperm from low fertility bulls have either damaged DNA or an alteration of the protamine packing around the DNA. Data to support this line of reasoning is the finding that sperm DNA of low fertility bulls has an increased ability to denature upon acid exposure [44]. Low embryonic development in bulls with normal fertilization rates may also be caused by reciprocal translocations leading to embryonic death [45,46].
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137
To investigate possible paternal influences on embryonic apoptosis, two approaches may be used. The first is to use bulls with comparable sperm quality but different in vivo fertility. Such bulls may possess intrinsic sperm defects that become only obvious after fertilization. Using this approach we found that as far as apoptosis at the blastocyst stage is concerned, no differences could be detected between bulls of low and high fertility [47]. However, a paternal effect at earlier cleavage stages cannot be excluded. The second approach is to induce sperm defects in some way, and evaluate the effect of embryonic development. Sperm defects can be induced in vivo; acute scrotal heating is a model for male subfertility and leads to subfertile semen and reduced embryo development in mice [48] and bovine [49]. Paternal heat stress causes embryonic arrest at the zygote stage in mice due to damage to nuclear remodeling and pronuclear formation—may inhibit the first tour of DNA replication and may impair the regulatory balance between cell death genes and cell survival genes. Heat stress has been shown to damage the chromatin structure of the sperm in the epididymis of the mouse and the bull, and increased the number of abnormal spermatozoa, which give rise to embryos with higher levels of caspase activity [50]. Another way to damage sperm is to irradiate them with X- or gamma rays after ejaculation [51]. The DNA damaged spermatozoa did not show morphological abnormalities or decreased motility. They were capable of fertilizing oocytes, but development was completely blocked due to initiation of apoptosis after the second or third cleavage, after the maternal-zygotic transition. In this case, repair of the paternal DNA damage by the oocyte was not possible and it was later destroyed by the apoptotic machinery of the embryo proper [51]. 3.2. Spermatozoa fit to sustain embryo development It is very difficult to select those spermatozoa, which are most fit to fertilize and produce embryos, and numerous reviews have been written on the evaluation of sperm attributes which may be correlated with fertility [52]. Suffice it to say that sperm selection in vivo requires a lengthy traversing of the female genital tract, with a period of storage in the isthmus, followed by a sperm release from the oviductal epithelium towards the mature oocyte [53]. This process is mimicked in vitro by putting millions of sperm of different capacitation status next to hundreds of cumulus–oocyte complexes. However, the best approach we have until now to select sperm fit for
S135
fertilization is let the cumulus cells do it for us. Removal of the cumulus cells before fertilization significantly reduces fertilization rates [54], probably because the cumulus cells serve as sperm attractants [55]. The presence or absence of cumulus cells during fertilization does not affect the apoptotic cell ratio in the subsequently formed blastocysts [56]. In this way, cumulus cells can contribute to the quantity of embryos, which are being generated, but not to the quality. 4. Perspectives and conclusions Gametic origin definitely has an influence on early embryo development. No matter how hard we try to select only the fit oocytes and spermatozoa for procedures involving in vitro embryo production, nature is always one step ahead of us. With invasive techniques we can already determine some aspects, which matter for embryonic development, such as normal undamaged DNA for the spermatozoa, and (large) oocytes with a sufficient stockpile of mRNA and proteins, which are derived from a healthy follicular environment. Cumulus cells may enlarge positive or negative effects from the maturation environment. Conscientious selection of sperm donors, of oocytes and of culture media is therefore imperative when embryos need to be produced for transfer purposes, to warrant optimal embryo quality. References [1] Betteridge KJ. Phylogeny, ontogeny and embryo transfer. Theriogenology 1995;44:1061–98. [2] Baker MA, Aitken RJ. Reactive oxygen species in spermatozoa: methods for monitoring and significance for the origins of genetic disease and infertility. Reprod Biol Endocrinol 2005;3:67. [3] Leese HJ, Donnay I, Thompson JG. Human assisted conception: a cautionary tale. Lessons from domestic animals. Hum Reprod 1998;13:184–202. [4] Reynaud K, Driancourt MA. Oocyte attrition. Mol Cell Endocrinol 2000;163:101–8. [5] Hussein TS, Froiland DA, Amato F, Thompson JG, Gilchrist RB. Oocytes prevent cumulus cell apoptosis by maintaining a morphogenic paracrine gradient of bone morphogenetic proteins. J Cell Sci 2005;118:5257–68. [6] Yuan YQ, Peelman L, Williams JL, Van Zeveren A, de Kruif A, Law A, et al. Mapping and transcription profiling of CASP1, 3, 6, 7, and 8 in relation to caspase activity in the bovine cumulus– oocyte complex. Anim Genet 2004;35:234–7. [7] Papandile A, Tyas D, O’Malley DM, Warner CM. Analysis of caspase-3, caspase-8 and caspase-9 activities in mouse oocytes and zygotes. Zygote 2004;12:57–64. [8] Brevini TAL, Cillo F, Antonini S, Tosetti V, Gandolfi F. Temporal and spatial control of gene expression in early embryos of farm animals. Reprod Fertil Dev 2007;19:35–42.
S136
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137
[9] Wang Q, Sun Q-Y. Evaluation of oocyte quality: morphological, cellular and molecular predictors. Reprod Fertil Dev 2007;19:1– 12. [10] Leroy JLMR, Vanholder T, Mateusen B, Christophe A, Opsomer G, de Kruif A, et al. Non-esterified fatty acids in follicular fluid of dairy cows and their effect on developmental capacity of bovine oocytes in vitro. Reproduction 2005;130:485–95. [11] Webb R, Garnsworthy PC, Gong J-G, Armstrong D. Control of follicular growth: local interactions and nutritional influences. J Anim Sci 2004;82:E63–74. [12] Lequarre´ AS, Vigneron C, Ribaucour F, Holm P, Donny I, Dalbies-Tran R, et al. Influence of antral follicle size on oocyte characteristics and embryo development in the bovine. Theriogenology 2005;63:841–59. [13] Fair T, Hyttel P, Greve T. Bovine oocyte diameter in relation to maturational competence and transcriptional activity. Mol Reprod Dev 1995;42:437–42. [14] Hyttel P, Viuff D, Fair T, Laurincik J, Thomsen PD, Callesen H, et al. Ribosomal RNA gene expression and chromosome aberrations in bovine oocytes and preimplantation embryos. Reproduction 2001;122:21–30. [15] Vandaele L, Mateusen B, Maes D, de Kruif A, Van Soom A. Temporal detection of caspase-3 and -7 in bovine in vitro produced embryos of different developmental capacity. Reproduction 2007;133:709–18. [16] Jurisicova A, Acton BM. Deadly decisions: the role of genes regulating programmed cell death in human preimplantation embryo development. Reproduction 2004;128:281–91. [17] Han Z, Chung YG, Gao S, Latham KE. Maternal factors controlling blastomere fragmentation in early mouse embryos. Biol Reprod 2005;72:612–8. [18] Anguita B, Vandaele L, Mateusen B, Maes D, Van Soom A. Developmental competence of bovine oocytes is not related to apoptosis incidence in oocytes, cumulus cells and blastocysts. Theriogenology 2006;673:537–49. [19] Hendriksen PJ, Vos PL, Steenweg WN, Bevers MM, Dieleman SJ. Bovine follicular development and its effect on the in vitro competence of oocytes. Theriogenology 2000;53:11–20. [20] Yang MY, Rajamahendran R. Morphological and biochemical identification of apoptosis in small, medium, and large bovine follicles and the effects of follicle-stimulating hormone and insulin-like growth factor-I on spontaneous apoptosis in cultured bovine granulosa cells. Biol Reprod 2000;62:1209–17. [21] Sirard MA, Richard F, Blondin P, Robert C. Contribution of the oocyte to embryo quality. Theriogenology 2006;65:126–36. [22] Ikeda S, Imai H, Yamada M. Apoptosis in cumulus cells during in vitro maturation of bovine cumulus-enclosed oocytes. Reproduction 2003;125:369–76. [23] Lee KS, Joo BS, Na YJ, Yoon MS, Choi OH, Kim WW. Cumulus cells apoptosis as an indicator to predict the quality of oocytes and the outcome of IVF-ET. J Assist Reprod Genet 2001;18:490– 8. [24] Yuan YQ, Van Soom A, Leroy JLMR, Dewulf J, Van Zeveren A, de Kruif A, et al. Apoptosis in cumulus cells, but not in oocytes, may influence bovine embryonic developmental competence. Theriogenology 2005;63:2147–63. [25] Tanghe S, Van Soom A, Nauwynck H, Coryn M, de Kruif A. Functions of the cumulus oophorus during oocyte maturation, ovulation and fertilization. Mol Reprod Dev 2002;61:414–24. [26] Ikeda S, Saeki K, Imai H, Yamada M. Abilities of cumulus and granulosa cells to enhance the developmental competence of bovine oocytes during in vitro maturation period are promoted
[27]
[28]
[29]
[30]
[31]
[32]
[33]
[34]
[35]
[36]
[37]
[38]
[39] [40] [41]
[42]
[43]
by midkine; a possible implication of its apoptosis suppressing effects. Reproduction 2006;132:549–57. Mourot M, Dufort I, Gravel C, Algriany O, Dieleman S, Sirard M-A. The influence of follicle size, FSH-enriched maturation medium, and early cleavage on bovine oocyte maternal mRNA levels. Mol Reprod Dev 2006;73:1367–79. Avery B, Strobech L, Jacobsen T, Bogh IB, Greve T. In vitro maturation of bovine cumulus–oocyte complexes in undiluted follicular fluid: effect on nuclear maturation, pronucleus formation and embryo development. Theriogenology 2003;59:987–99. Bijttebier J, Van Soom A, Mateusen B, Maes D. In vitro maturation of porcine oocytes in medium supplemented with ultra-filtered porcine follicular fluid. Reprod Dom Anim 2006;41:306. Nicholas B, Alberio R, Fouladi-Nashta AA, Webb R. Relationship between low-molecular-weight insulin-like growth factorbinding proteins, caspase-3 activity, and oocyte quality. Biol Reprod 2005;72:796–804. Van Soom A, Van Vlaenderen I, Mahmoudzadeh AR, Deluyker H, de Kruif A. Compaction rate of in vitro fertilized bovine embryos related to the interval from insemination to first cleavage. Theriogenology 1992;38:905–19. Fair T, Murphy M, Rizos D, Moss C, Martin F, Boland MP, et al. Analysis of differential maternal mRNA expression in developmentally competent and incompetent bovine two-cell embryos. Mol Reprod Dev 2004;67:136–44. Dode MA, Dufort I, Massicotte L, Sirard MA. Quantitative expression of candidate genes for developmental competence in bovine two-cell embryos. Mol Reprod Dev 2006;73:288–97. Donnison M, Pfeffer PL. Isolation of genes associated with developmentally competent bovine oocytes and quantitation of their levels during development. Biol Reprod 2004;71: 1813–21. Van Soom A, Ysebaert MT, de Kruif A. Relationship between timing of development, morula morphology, and cell allocation to inner cell mass and trophectoderm in in vitro-produced bovine embryos. Mol Reprod Dev 1997;47:47–56. Short RV. The testis: the witness of the mating system, the site of mutation and the engine of desire. Acta Paediatr 1997;422 (Suppl):3–7. Coetzee K, Kruge TF, Lombard CJ. Predictive value of normal sperm morphology: a structured literature review. Hum Reprod Update 1998;4:73–82. Sutovsky P, Van Leyen K, McCaulet T, Day BN, Sutovsky M. Degradation of paternal mitochondria after fertilization: implications for heteroplasmy, assisted reproductive technologies and mtDNA inheritance. Reprod Biomed Online 2004;8:24– 33. Aitken RJ, Krausz C. Oxidative stress, DNA damage and the Y chromosome. Reproduction 2001;122:497–506. Jaroudi S, SenGupta S. DNA repair in mammalian embryos. Mutat Res 2007;635:53–77. Menezo YJ. Paternal and maternal factors in preimplantation embryogenesis: interaction with the biochemical environment. Reprod Biomed Online 2006;12:616–21. Ji BT, Shu XO, Linet MS, Zhengf W, Wacholder S, Gao YT, et al. Paternal cigarette smoking and the risk of childhood cancer among offspring of nonsmoking mothers. J Natl Cancer Inst 1997;89:238–44. Eid LN, Lorton SP, Parrish JJ. Paternal influence on S-phase in the first cell cycle of the bovine embryo. Biol Reprod 2004;51:1232–7.
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137 [44] Ballachey BE, Evenson DP, Saacke RG. The sperm chromatin structure assay. Relationship with alternate tests of semen quality and heterospermic performance of bulls. J Androl 1988;9: 109–15. [45] Andersson M, Aalto J, Gustavsson I. Embryo quality and andrological study of two subfertile bulls versus five control bulls with normal fertility. Theriogenology 1992;38:623–31. [46] Ducos A, Dumont P, Seguela A, Pinton A, Berland H, BrunBaronnat C, et al. A new reciprocal translocation in a subfertile bull. Genet Sel Evol 2000;32:589–98. [47] Vandaele L, Mateusen B, Maes D, de Kruif A, Van Soom A. Is apoptosis in bovine in vitro produced embryos related to early developmental kinetics and in vivo bull fertility? Theriogenology 2006;65:1691–703. [48] Zhu B-K, Setchell BP. Effects of paternal heat stress on the in vivo development of preimplantation embryos in the mouse. Reprod Nutr Dev 2004;44:617–29. [49] Walters AH, Saacke RG, Pearson RE, Gwazdauskas FC. Assessment of pronuclear formation following in vitro fertilization with bovine spermatozoa obtained after thermal insulation of the testis. Theriogenology 2006;65:1016–28. [50] Walters AH, Saacke RG, Pearson RE, Gwazdauskas FC. The incidence of apoptosis after IVF with morphologically
[51]
[52] [53]
[54]
[55]
[56]
S137
abnormal bovine spermatozoa. Theriogenoloy 2005;64: 1404–21. Fatehi AN, Bevers MM, Schoevers E, Roelen BAJ, Colenbrander B, Gadella BM. DNA damage in bovine sperm does not block fertilization and early embryonic development but induces apoptosis after the first cleavages. J Androl 2006;27: 176–88. Rodriguez-Martinez H. State of the art in farm animal sperm evaluation. Reprod Fertil Dev 2007;19:91–102. Suarez SS. Interactions of spermatozoa with the female reproductive tract: inspiration for assisted reproduction. Reprod Fertil Dev 2007;19:91–102. Tanghe S, Van Soom A, Mehrzad J, Maes D, Duchateau P, de Kruif A. Cumulus contributions during bovine fertilization in vitro. Theriogenology 2003;60:135–49. Van Soom A, Tanghe S, De Pauw I, Maes D, de Kruif A. Function of the cumulus oophorus before and during mammalian fertilization. Reprod Dom Anim 2002;37:141–51. Fatehi AN, Roelen BA, Colenbrander B, Schoevers EJ, Gadella BM, Bevers MM, et al. Presence of cumulus cells during in vitro fertilization protects the bovine oocyte against oxidative stress and improves first cleavage but does not affect further development. Zygote 2005;13:177–85.
Gamete origin in relation to early embryo development A. Van Soom a,*, L. Vandaele a, K. Goossens b, A. de Kruif a, L. Peelman b a
Reproductive Biology Unit, Department of Reproduction, Obstetrics and Herd Health, Faculty of Veterinary Medicine, Ghent University, Salisburylaan 133, B-9820 Merelbeke, Belgium b Molecular Biology Unit, Heidestraat 19, Faculty of Veterinary Medicine, 9820 Merelbeke, Belgium
Abstract Fertilization in vivo requires a complex series of selection events to occur in order to guarantee that only the fittest gametes take part in the fusion process and give rise to a viable embryo. Conventional practice in bovine in vitro fertilization however is to select oocytes and sperm by quite crude procedures. It is therefore not inconceivable that essentially unfit gametes may drive aberrant embryo development in vitro. Abnormal embryonic cells are being removed by apoptosis, which is a physiological process in embryos. Only an excess or a lack of apoptosis can lead to embryonic death or abnormal development. Suboptimal culture conditions undoubtedly contribute to undue embryonic apoptosis, but the intrinsic quality of the oocyte may also be a causative factor. It is generally accepted that the oocyte is in control of early embryogenesis, but is it also in control of future embryonic suicide? Is a compromised follicular environment predestining the oocyte to a dire fate? What is the contribution of the cumulus cells to oocyte quality, and can they rescue it from early demise? And what can be said about the origin of the spermatozoa? Research in human in vitro fertilization has definitely shown that factors such as paternal age, smoking and other sperm stressors can contribute to abnormal embryo development and even diseased offspring. This review will address the questions raised above, and will describe what is known about the cellular and molecular biology that may account for abnormal bovine embryo development caused by gamete origin. # 2007 Elsevier Inc. All rights reserved. Keywords: Gamete origin; Embryo; Apoptosis
1. Introduction A tantalizing phrase which has been introduced for the first time in Darwin’s Origin of species is ‘‘survival of the fittest’’. Since the title of this talk reminded me in some way to evolutionary biology, I wondered whether this very concept could not be used to explain embryonic development as well. In modern biology, the term fitness measures reproductive success and is not explicit about the specific ways in which organisms (or in our case, gametes or cells) can be ‘‘fit’’ as in
* Corresponding author. Tel.: +32 9 264 7561; fax: +32 9 264 7797. E-mail address: [email protected] (A. Van Soom). 0093-691X/$ – see front matter # 2007 Elsevier Inc. All rights reserved. doi:10.1016/j.theriogenology.2007.03.019
‘‘having phenotypic characteristics which enhance survival and reproduction’’. The methods we use to select the gametes meant for assisted reproduction and the subsequent treatments we inflict on them are surely imposing a kind of selection pressure, albeit not a natural but merely an artificial one. Natural selection is the central concept of Darwinian theory – the fittest survive, and can give rise to offspring – but do we actually select the fittest gametes? Which are the fittest? And how is ‘‘fitness’’ defined? The study of links between development and evolution has been relatively neglected by animal scientists and veterinarians, who prefer to focus their scientific concepts on the animal as a whole, or to find potential applications for a certain technique [1]. The applied embryology that we are
S132
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137
practicing does not aim to analyze evolutionary mechanisms, since there is no reason to believe that these mechanisms are involved in the process of embryo development in vitro. But can we be sure that this is really so? Are not there numerous examples in assisted reproductive biology to prove that deliberate selection of gametes and/or embryos is causing an effect on the actual outcome? We probably all agree that embryo development is being affected by gamete origin. Gamete origin can be defined at different levels: it can relate to the donor, to the in vivo or in vitro maturation status of the gamete, and to its supporting environment. When these determinants of gamete origin are inferior, they will predestine the resulting embryo for an ill-fated future. Such future can be on short-term aberrant fertilization and abnormal embryo development, resulting in developmental block, retarded development or embryonic demise, or, on the long term, lack of implantation and diseased offspring [2,3]. In this review, we will focus on how we currently define which gametes are fit to take part in the process of reproduction, what the impact of their origin may have on subsequent early embryonic development, and what role apoptosis or programmed cell death plays in determining the fate of the embryo. We hypothesize that the basic machinery to cause embryonic apoptosis may already be present in the oocyte or the sperm. We will review data from the literature and present experimental evidence, obtained in our laboratory mainly in cattle, to support this hypothesis. 2. Of follicles, oocytes and apoptosis Many more follicles are present in the mammalian ovary than will eventually ovulate [4]. Solid evidence is available that the massive loss of germ cells during oogenesis is caused by oocyte apoptosis. Therefore, the normal fate for female germ cells during oogenesis is death. Interestingly, once an antrum has been formed, follicular atresia is mainly driven by apoptosis of granulosa cells, and the oocyte itself is quite resistant to cell death [4]. In fact it can even protect the surrounding cumulus cells against apoptosis [5]. This does not mean, however, that the oocyte is not capable of initiating apoptosis. Transcripts of caspases-3, -7, -8 and -9 have been demonstrated in bovine [6] and mouse oocytes [7], showing that unfertilized oocytes have the machinery to undergo apoptosis by using either the extrinsic (caspase-8 dependent) or intrinsic (caspase-9 dependent) pathways. The different balance in expression of pro- and anti-apoptotic genes may shift the oocyte’s
developmental potential towards either cell death or cell survival, and may be a crucial factor in determining the quality of the oocyte. But how can we predict which oocyte will give rise to a viable embryo and which to a failing one, and can we influence the fitness of the oocyte by means of better maturation conditions? Several aspects of oocyte origin can influence eventual fertilization outcome. During follicular development, oocytes and follicles are forming functional units. This is especially true if we look at the cumulus–oocyte interactions during the late stages of ovogenesis, including oocyte capacitation and maturation. Whereas the intrinsic quality of an oocyte is determined by the presence of a normal set of chromosomes and by its mitochondrial genome, the microenvironment of the oocyte, be it either the preovulatory follicle or the in vitro maturation conditions can influence, on the other hand, the fate of the transcripts and proteins accumulated in the oocyte [8–10]. 2.1. Oocytes unfit to sustain embryo development Are we at risk to select unfit oocytes, when we do apply the morphological criteria which most scientists use very rigorously? The answer is yes. It has long been accepted that the population of oocytes which is being used for in vitro embryo production is a very heterogeneous one. Immature oocytes are being aspirated from antral follicles which are typically between 2 and 8 mm in diameter [11]. A lower developmental potential has already been demonstrated for oocytes derived from smaller follicles (<4 mm) versus those from larger follicles (>6 mm) although no differences could be detected directly at the oocyte level, as far as global profile of transcripts, the pattern of protein neosynthesis and the kinetics of meiosis resumption was concerned [12]. However, it has been demonstrated that bovine follicles smaller than 3 mm contain oocytes, which are smaller than 110 mm [13]. Reaching a critical diameter of 110 mm coincides, for the bovine oocyte at least, with the acquisition of full competence for completing maturation to metaphase II [14]. Embryos generated by smaller oocytes (<110 mm) displayed a higher incidence of apoptosis throughout embryo development as assessed by fluorescent detection of active caspase-3 and -7 in cleaved embryos and morulae [15]. This effect was present until 117 hpi, and may indicate that such embryos are derived from oocytes, which do not possess the complete protein synthetic machinery to prevent embryonic apoptosis. About one third of bovine oocytes possess the transcripts for caspases, but active caspases
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137
are not detectable yet [6]. If however inactive caspases are activated in unfit oocytes, fertilization of such oocytes may lead to embryo fragmentation and impaired embryonic development, because the oocyte cytoplasm cannot provide the necessary transcripts and proteins for adequate suppression of apoptotic pathways [16,17]. Evidence to support this is provided by the finding that oocytes smaller than 110 mm showed in 80% of the cases DNA fragmentation before maturation, whereas this was only 20% for oocytes of larger diameters [18]. More evidence for the importance of follicular origin in determining embryonic fate, is the finding that larger antral follicles are required (>6–8 mm) for collection of oocytes which have the competence for both nuclear and cytoplasmic maturation [19]. Such follicles can be either atretic or healthy in cattle: many more apoptotic granulosa cells are present in the atretic follicles than in the healthy ones, whilst no apoptotic cells could be demonstrated in the cumulus–oocyte complexes from the same follicles [20]. Matters are complicated by the fact that oocytes from early atretic follicles display increased developmental competence [21], and oocyte removal from the follicular environment may artificially induce cumulus cell apoptosis, since cumulus–oocyte complexes show a spontaneous onset of apoptosis during in vitro maturation [20,22], or during prolonged manipulation, predominantly in the outer layers of the cumulus cells (Anguita, unpublished results). How important are these apoptotic cumulus cells for the fate of the oocyte? It has been argued that a high prevalence of apoptotic cumulus cells surrounding the oocyte may be predictive for low developmental potential, in human [23] and in cattle studies [24]. Cumulus cells communicate with each other and with the oocyte via gap junctions (reviewed by [25]), which are essential for transport of nutrients and regulatory factors, but these contacts are lost during the process of maturation. Anyhow, components of follicular fluid, which have been shown to be detrimental for oocyte developmental capacity, such as palmitic and stearic acid, caused increased levels of apoptosis in cumulus cells [10], which is indicative for a causal relationship. On the other hand, growth factors, such as midkine, which have been shown to be beneficial to oocyte developmental competence, have an anti-apoptotic effect on cumulus cells and can exert their effect even through isolated cumulus cells [26]. 2.2. Oocytes fit to sustain embryo development A second maybe much more important question we must ask ourselves is which oocytes are actually fit for
S133
normal embryo production. This question is far more difficult to answer, because no reliable non-invasive methods for oocyte selection are at present available, besides the detection of glucose-6-phosphate dehydrogenase by means of brilliant cresyl blue staining, as recently been reviewed by Ref. [9]. They also stated that analysis of factors in follicular fluid/cells may hold promise to foretell oocyte quality on an individual basis. Probably the most promising approaches in this field are the use of functional genomics, transcriptomics and proteomics, since fit oocytes need to contain a sufficient stockpile of mRNA and proteins to allow embryo development to occur until after embryonic genome activation. First of all, fully-grown oocytes from larger follicles are definitely better equipped to generate embryos. The higher embryo yield of oocytes from larger follicles has been related directly to differences in mRNA transcripts in a recent study [27]. By using the powerful approach of suppressive subtractive hybridization, it was demonstrated that several genes are differentially expressed according to different follicular sizes. Genes involved in cell cycle regulation such as cyclin dependent kinase subunit 1 (CKS1B), pituitary tumor-transforming 1 (PTTG1) and cyclin B2 (CCNB2), and histone 2A (H2A) which contributes to the chromatin support in the early embryo, are gradually upgraded according to follicular size [27]. Second, it would be quite logical to assume that preovulatory follicular fluid provides the best environment for the maturing oocyte. For bovine oocytes, undiluted follicular fluid is not the optimal maturation environment [28], whereas it is added as a routine during pig oocyte maturation. We have recently conducted a number of experiments to get a better understanding of the role of follicular fluid on pig oocyte maturation and found that components larger than 10 kDa have a positive effect on nuclear maturation, but were not able to show an effect on cytoplasmic maturation [29]. Preovulatory porcine follicular fluid however was superior over autologous serum to sustain pig cytoplasmic maturation (Bijttebier, unpublished results). Further studies using proteomic analysis are in progress to identify the molecules involved. Recent data in cattle have also shown that low IGFBP-levels in follicular fluid can be used to select fit oocytes [30]. It remains to be determined whether addition of these molecules to maturation medium can rescue oocytes from early demise. A third determinant of oocyte competence is early cleavage [31,32]. One approach is to use transcriptomics to differentiate between fast and slow cleavers,
S134
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137
and again higher levels of histone H2A [33] and histone H3A [32] have been detected in embryos derived from fit oocytes, which is logical because histones are needed to replace the protamines of the sperm nucleus after fertilization and to assemble embryonic DNA into chromatin for the first cleavages. Other transcripts which were differentially expressed in fast cleavers were stem-loop binding protein (SLBP), which is involved in histone stabilisation [34], YEAF, which is a transcriptional repressor, and isocitrate dehydrogenase (IDH), which modulates oxidative damage [33]. Another approach, which we have used repeatedly in the past, was to look at quality of embryos derived from either fit (fast cleavers) or unfit (slow) oocytes. We found that fast cleavers had higher chances to become a morula or blastocyst, and to display an excellent morphology [31,35]. Moreover, fast cleavers had significantly lower apoptotic indices at 45, 80 and 117 hpi, indicating a maternal effect [15]. The lower incidence of apoptosis may be attributed to the superior stockpile of mRNAs and proteins, which typecast such fit oocytes, but may also be influenced by mitochondria present in fit oocytes. It is generally accepted that the oocyte is in control of early embryogenesis, and our data confirm that it is at least partially in control of future embryonic suicide. Selection of fit oocytes must be followed by optimal insemination and culture conditions to prevent apoptosis to occur. 3. Of spermatozoa, DNA damage and fertilization It has been said that the engine of evolution is the testis [36], with its continuous production of large numbers of spermatozoa by a spermatogenic process that has no means of selecting against phenotypic or genotypic abnormalities. Nature, however, prevents the procreation of severe mutations in the male germ line by selecting against structurally deviant spermatozoa, which have a lower chance to reach and fertilize a mature oocyte [37]. In addition, the sperm mitochondria, which may also carry mutations, are destroyed in the oocyte after fertilization [38]. Two types of DNA damage are however typical for the male germ line: replication errors and DNA fragmentation [39]. Replication errors are more prevalent in the spermatozoa of older men and have been linked with the occurrence of dominant genetic disease, but may be less important for domestic animals, whose lifespan is shorter. DNA damage on the contrary, may occur frequently in ejaculates of otherwise healthy animals, and may arise from three sources: (1) oxidative
stress, (2) abortive Fas-mediated apoptosis and (3) deficiencies in chromatin packaging leading to DNA strand breaks [39]. Fortunately, DNA repair mechanisms which rely on the maternal mRNAs and proteins stored in the oocyte are able to mend errors in DNA, leading to healthy offspring [40]. If an oocyte is fertilized with DNA-damaged spermatozoa, G1-associated DNA repair mechanisms become activated, leading to a dramatic suppression of pronuclear DNA-synthesis via a p53dependent mechanism. Mice lacking such DNA repair enzymes are not viable with preimplantation embryonic death as a result [40]. Which characteristics are needed for spermatozoa to sustain embryo development? It is has been demonstrated that bulls can differ in embryo generating capacity, even if they display similar fertilization rates in vitro. In the early years of human in vitro fertilization, it was postulated that sperm quality and early embryonic development were independent, but later on this was reviewed in favour of a paternal effect on early embryonic viability and further development [41]. Now there is a consensus that suboptimal spermatozoa have a negative influence on the integrity and the quality of the paternal DNA, and may lead to defective embryonic development or diseased offspring. For instance, failure of Sertoli cells to remove sufficient residual cytoplasm from the spermatozoa during spermatogenesis enhances free radical production in the ejaculates of infertile human patients [39]. Evidence exists that children from heavy smokers, whose semen is characterized by oxidative DNA base damage, are four to five times more likely to develop childhood cancer than children from non-smokers [42]. 3.1. Spermatozoa unfit to sustain embryo development Differences in development between embryos sired by different bulls can first be detected before cleavage to the two-cell stage (but after pronuclear formation) and thus before expression of the embryonic genome. Zygotes sired by low fertility bulls showed a delay in Sphase initiation compared to zygotes sired by highfertility bulls [43]: it has been hypothesized that sperm from low fertility bulls have either damaged DNA or an alteration of the protamine packing around the DNA. Data to support this line of reasoning is the finding that sperm DNA of low fertility bulls has an increased ability to denature upon acid exposure [44]. Low embryonic development in bulls with normal fertilization rates may also be caused by reciprocal translocations leading to embryonic death [45,46].
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137
To investigate possible paternal influences on embryonic apoptosis, two approaches may be used. The first is to use bulls with comparable sperm quality but different in vivo fertility. Such bulls may possess intrinsic sperm defects that become only obvious after fertilization. Using this approach we found that as far as apoptosis at the blastocyst stage is concerned, no differences could be detected between bulls of low and high fertility [47]. However, a paternal effect at earlier cleavage stages cannot be excluded. The second approach is to induce sperm defects in some way, and evaluate the effect of embryonic development. Sperm defects can be induced in vivo; acute scrotal heating is a model for male subfertility and leads to subfertile semen and reduced embryo development in mice [48] and bovine [49]. Paternal heat stress causes embryonic arrest at the zygote stage in mice due to damage to nuclear remodeling and pronuclear formation—may inhibit the first tour of DNA replication and may impair the regulatory balance between cell death genes and cell survival genes. Heat stress has been shown to damage the chromatin structure of the sperm in the epididymis of the mouse and the bull, and increased the number of abnormal spermatozoa, which give rise to embryos with higher levels of caspase activity [50]. Another way to damage sperm is to irradiate them with X- or gamma rays after ejaculation [51]. The DNA damaged spermatozoa did not show morphological abnormalities or decreased motility. They were capable of fertilizing oocytes, but development was completely blocked due to initiation of apoptosis after the second or third cleavage, after the maternal-zygotic transition. In this case, repair of the paternal DNA damage by the oocyte was not possible and it was later destroyed by the apoptotic machinery of the embryo proper [51]. 3.2. Spermatozoa fit to sustain embryo development It is very difficult to select those spermatozoa, which are most fit to fertilize and produce embryos, and numerous reviews have been written on the evaluation of sperm attributes which may be correlated with fertility [52]. Suffice it to say that sperm selection in vivo requires a lengthy traversing of the female genital tract, with a period of storage in the isthmus, followed by a sperm release from the oviductal epithelium towards the mature oocyte [53]. This process is mimicked in vitro by putting millions of sperm of different capacitation status next to hundreds of cumulus–oocyte complexes. However, the best approach we have until now to select sperm fit for
S135
fertilization is let the cumulus cells do it for us. Removal of the cumulus cells before fertilization significantly reduces fertilization rates [54], probably because the cumulus cells serve as sperm attractants [55]. The presence or absence of cumulus cells during fertilization does not affect the apoptotic cell ratio in the subsequently formed blastocysts [56]. In this way, cumulus cells can contribute to the quantity of embryos, which are being generated, but not to the quality. 4. Perspectives and conclusions Gametic origin definitely has an influence on early embryo development. No matter how hard we try to select only the fit oocytes and spermatozoa for procedures involving in vitro embryo production, nature is always one step ahead of us. With invasive techniques we can already determine some aspects, which matter for embryonic development, such as normal undamaged DNA for the spermatozoa, and (large) oocytes with a sufficient stockpile of mRNA and proteins, which are derived from a healthy follicular environment. Cumulus cells may enlarge positive or negative effects from the maturation environment. Conscientious selection of sperm donors, of oocytes and of culture media is therefore imperative when embryos need to be produced for transfer purposes, to warrant optimal embryo quality. References [1] Betteridge KJ. Phylogeny, ontogeny and embryo transfer. Theriogenology 1995;44:1061–98. [2] Baker MA, Aitken RJ. Reactive oxygen species in spermatozoa: methods for monitoring and significance for the origins of genetic disease and infertility. Reprod Biol Endocrinol 2005;3:67. [3] Leese HJ, Donnay I, Thompson JG. Human assisted conception: a cautionary tale. Lessons from domestic animals. Hum Reprod 1998;13:184–202. [4] Reynaud K, Driancourt MA. Oocyte attrition. Mol Cell Endocrinol 2000;163:101–8. [5] Hussein TS, Froiland DA, Amato F, Thompson JG, Gilchrist RB. Oocytes prevent cumulus cell apoptosis by maintaining a morphogenic paracrine gradient of bone morphogenetic proteins. J Cell Sci 2005;118:5257–68. [6] Yuan YQ, Peelman L, Williams JL, Van Zeveren A, de Kruif A, Law A, et al. Mapping and transcription profiling of CASP1, 3, 6, 7, and 8 in relation to caspase activity in the bovine cumulus– oocyte complex. Anim Genet 2004;35:234–7. [7] Papandile A, Tyas D, O’Malley DM, Warner CM. Analysis of caspase-3, caspase-8 and caspase-9 activities in mouse oocytes and zygotes. Zygote 2004;12:57–64. [8] Brevini TAL, Cillo F, Antonini S, Tosetti V, Gandolfi F. Temporal and spatial control of gene expression in early embryos of farm animals. Reprod Fertil Dev 2007;19:35–42.
S136
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137
[9] Wang Q, Sun Q-Y. Evaluation of oocyte quality: morphological, cellular and molecular predictors. Reprod Fertil Dev 2007;19:1– 12. [10] Leroy JLMR, Vanholder T, Mateusen B, Christophe A, Opsomer G, de Kruif A, et al. Non-esterified fatty acids in follicular fluid of dairy cows and their effect on developmental capacity of bovine oocytes in vitro. Reproduction 2005;130:485–95. [11] Webb R, Garnsworthy PC, Gong J-G, Armstrong D. Control of follicular growth: local interactions and nutritional influences. J Anim Sci 2004;82:E63–74. [12] Lequarre´ AS, Vigneron C, Ribaucour F, Holm P, Donny I, Dalbies-Tran R, et al. Influence of antral follicle size on oocyte characteristics and embryo development in the bovine. Theriogenology 2005;63:841–59. [13] Fair T, Hyttel P, Greve T. Bovine oocyte diameter in relation to maturational competence and transcriptional activity. Mol Reprod Dev 1995;42:437–42. [14] Hyttel P, Viuff D, Fair T, Laurincik J, Thomsen PD, Callesen H, et al. Ribosomal RNA gene expression and chromosome aberrations in bovine oocytes and preimplantation embryos. Reproduction 2001;122:21–30. [15] Vandaele L, Mateusen B, Maes D, de Kruif A, Van Soom A. Temporal detection of caspase-3 and -7 in bovine in vitro produced embryos of different developmental capacity. Reproduction 2007;133:709–18. [16] Jurisicova A, Acton BM. Deadly decisions: the role of genes regulating programmed cell death in human preimplantation embryo development. Reproduction 2004;128:281–91. [17] Han Z, Chung YG, Gao S, Latham KE. Maternal factors controlling blastomere fragmentation in early mouse embryos. Biol Reprod 2005;72:612–8. [18] Anguita B, Vandaele L, Mateusen B, Maes D, Van Soom A. Developmental competence of bovine oocytes is not related to apoptosis incidence in oocytes, cumulus cells and blastocysts. Theriogenology 2006;673:537–49. [19] Hendriksen PJ, Vos PL, Steenweg WN, Bevers MM, Dieleman SJ. Bovine follicular development and its effect on the in vitro competence of oocytes. Theriogenology 2000;53:11–20. [20] Yang MY, Rajamahendran R. Morphological and biochemical identification of apoptosis in small, medium, and large bovine follicles and the effects of follicle-stimulating hormone and insulin-like growth factor-I on spontaneous apoptosis in cultured bovine granulosa cells. Biol Reprod 2000;62:1209–17. [21] Sirard MA, Richard F, Blondin P, Robert C. Contribution of the oocyte to embryo quality. Theriogenology 2006;65:126–36. [22] Ikeda S, Imai H, Yamada M. Apoptosis in cumulus cells during in vitro maturation of bovine cumulus-enclosed oocytes. Reproduction 2003;125:369–76. [23] Lee KS, Joo BS, Na YJ, Yoon MS, Choi OH, Kim WW. Cumulus cells apoptosis as an indicator to predict the quality of oocytes and the outcome of IVF-ET. J Assist Reprod Genet 2001;18:490– 8. [24] Yuan YQ, Van Soom A, Leroy JLMR, Dewulf J, Van Zeveren A, de Kruif A, et al. Apoptosis in cumulus cells, but not in oocytes, may influence bovine embryonic developmental competence. Theriogenology 2005;63:2147–63. [25] Tanghe S, Van Soom A, Nauwynck H, Coryn M, de Kruif A. Functions of the cumulus oophorus during oocyte maturation, ovulation and fertilization. Mol Reprod Dev 2002;61:414–24. [26] Ikeda S, Saeki K, Imai H, Yamada M. Abilities of cumulus and granulosa cells to enhance the developmental competence of bovine oocytes during in vitro maturation period are promoted
[27]
[28]
[29]
[30]
[31]
[32]
[33]
[34]
[35]
[36]
[37]
[38]
[39] [40] [41]
[42]
[43]
by midkine; a possible implication of its apoptosis suppressing effects. Reproduction 2006;132:549–57. Mourot M, Dufort I, Gravel C, Algriany O, Dieleman S, Sirard M-A. The influence of follicle size, FSH-enriched maturation medium, and early cleavage on bovine oocyte maternal mRNA levels. Mol Reprod Dev 2006;73:1367–79. Avery B, Strobech L, Jacobsen T, Bogh IB, Greve T. In vitro maturation of bovine cumulus–oocyte complexes in undiluted follicular fluid: effect on nuclear maturation, pronucleus formation and embryo development. Theriogenology 2003;59:987–99. Bijttebier J, Van Soom A, Mateusen B, Maes D. In vitro maturation of porcine oocytes in medium supplemented with ultra-filtered porcine follicular fluid. Reprod Dom Anim 2006;41:306. Nicholas B, Alberio R, Fouladi-Nashta AA, Webb R. Relationship between low-molecular-weight insulin-like growth factorbinding proteins, caspase-3 activity, and oocyte quality. Biol Reprod 2005;72:796–804. Van Soom A, Van Vlaenderen I, Mahmoudzadeh AR, Deluyker H, de Kruif A. Compaction rate of in vitro fertilized bovine embryos related to the interval from insemination to first cleavage. Theriogenology 1992;38:905–19. Fair T, Murphy M, Rizos D, Moss C, Martin F, Boland MP, et al. Analysis of differential maternal mRNA expression in developmentally competent and incompetent bovine two-cell embryos. Mol Reprod Dev 2004;67:136–44. Dode MA, Dufort I, Massicotte L, Sirard MA. Quantitative expression of candidate genes for developmental competence in bovine two-cell embryos. Mol Reprod Dev 2006;73:288–97. Donnison M, Pfeffer PL. Isolation of genes associated with developmentally competent bovine oocytes and quantitation of their levels during development. Biol Reprod 2004;71: 1813–21. Van Soom A, Ysebaert MT, de Kruif A. Relationship between timing of development, morula morphology, and cell allocation to inner cell mass and trophectoderm in in vitro-produced bovine embryos. Mol Reprod Dev 1997;47:47–56. Short RV. The testis: the witness of the mating system, the site of mutation and the engine of desire. Acta Paediatr 1997;422 (Suppl):3–7. Coetzee K, Kruge TF, Lombard CJ. Predictive value of normal sperm morphology: a structured literature review. Hum Reprod Update 1998;4:73–82. Sutovsky P, Van Leyen K, McCaulet T, Day BN, Sutovsky M. Degradation of paternal mitochondria after fertilization: implications for heteroplasmy, assisted reproductive technologies and mtDNA inheritance. Reprod Biomed Online 2004;8:24– 33. Aitken RJ, Krausz C. Oxidative stress, DNA damage and the Y chromosome. Reproduction 2001;122:497–506. Jaroudi S, SenGupta S. DNA repair in mammalian embryos. Mutat Res 2007;635:53–77. Menezo YJ. Paternal and maternal factors in preimplantation embryogenesis: interaction with the biochemical environment. Reprod Biomed Online 2006;12:616–21. Ji BT, Shu XO, Linet MS, Zhengf W, Wacholder S, Gao YT, et al. Paternal cigarette smoking and the risk of childhood cancer among offspring of nonsmoking mothers. J Natl Cancer Inst 1997;89:238–44. Eid LN, Lorton SP, Parrish JJ. Paternal influence on S-phase in the first cell cycle of the bovine embryo. Biol Reprod 2004;51:1232–7.
A. Van Soom et al. / Theriogenology 68S (2007) S131–S137 [44] Ballachey BE, Evenson DP, Saacke RG. The sperm chromatin structure assay. Relationship with alternate tests of semen quality and heterospermic performance of bulls. J Androl 1988;9: 109–15. [45] Andersson M, Aalto J, Gustavsson I. Embryo quality and andrological study of two subfertile bulls versus five control bulls with normal fertility. Theriogenology 1992;38:623–31. [46] Ducos A, Dumont P, Seguela A, Pinton A, Berland H, BrunBaronnat C, et al. A new reciprocal translocation in a subfertile bull. Genet Sel Evol 2000;32:589–98. [47] Vandaele L, Mateusen B, Maes D, de Kruif A, Van Soom A. Is apoptosis in bovine in vitro produced embryos related to early developmental kinetics and in vivo bull fertility? Theriogenology 2006;65:1691–703. [48] Zhu B-K, Setchell BP. Effects of paternal heat stress on the in vivo development of preimplantation embryos in the mouse. Reprod Nutr Dev 2004;44:617–29. [49] Walters AH, Saacke RG, Pearson RE, Gwazdauskas FC. Assessment of pronuclear formation following in vitro fertilization with bovine spermatozoa obtained after thermal insulation of the testis. Theriogenology 2006;65:1016–28. [50] Walters AH, Saacke RG, Pearson RE, Gwazdauskas FC. The incidence of apoptosis after IVF with morphologically
[51]
[52] [53]
[54]
[55]
[56]
S137
abnormal bovine spermatozoa. Theriogenoloy 2005;64: 1404–21. Fatehi AN, Bevers MM, Schoevers E, Roelen BAJ, Colenbrander B, Gadella BM. DNA damage in bovine sperm does not block fertilization and early embryonic development but induces apoptosis after the first cleavages. J Androl 2006;27: 176–88. Rodriguez-Martinez H. State of the art in farm animal sperm evaluation. Reprod Fertil Dev 2007;19:91–102. Suarez SS. Interactions of spermatozoa with the female reproductive tract: inspiration for assisted reproduction. Reprod Fertil Dev 2007;19:91–102. Tanghe S, Van Soom A, Mehrzad J, Maes D, Duchateau P, de Kruif A. Cumulus contributions during bovine fertilization in vitro. Theriogenology 2003;60:135–49. Van Soom A, Tanghe S, De Pauw I, Maes D, de Kruif A. Function of the cumulus oophorus before and during mammalian fertilization. Reprod Dom Anim 2002;37:141–51. Fatehi AN, Roelen BA, Colenbrander B, Schoevers EJ, Gadella BM, Bevers MM, et al. Presence of cumulus cells during in vitro fertilization protects the bovine oocyte against oxidative stress and improves first cleavage but does not affect further development. Zygote 2005;13:177–85.