Editor-in-Chief
Prof. Yi-Fei WANG,
Asian J Androl 2006; 8 (6): 643-673 |
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This web only provides the extract of this article. If you want to read the figures and tables, please reference the PDF full text on Blackwell Synergy. Thank you. - Review - Genetic and epigenetic risks of intracytoplasmic sperm injection method Ioannis Georgiou1, Maria Syrrou2, Nicolaos Pardalidis1, Konstantinos Karakitsios1, Themis Mantzavinos1, Nikolaos Giotitsas1, Dimitrios Loutradis3, Fotis Dimitriadis1, Motoaki Saito4, Ikuo Miyagawa3, Pavlos Tzoumis1, Anastasios Sylakos1, Nikolaos Kanakas1, Theodoros Moustakareas1, Dimitrios Baltogiannis1, Stavros Touloupides1, Dimitrios Giannakis1, Michael Fatouros1, Nikolaos Sofikitis1,3 1Laboratory of Molecular Urology and Genetics of Human Reproduction, Department of Urology, Ioannina University
School of Medicine, Ioannina 45110, Greece
Abstract Pregnancies achieved by assisted reproduction technologies, particularly by intracytoplasmic sperm injection (ICSI) procedures, are susceptible to genetic risks inherent to the male population treated with ICSI and additional risks inherent to this innovative procedure. The documented, as well as the theoretical, risks are discussed in the present review study. These risks mainly represent thatconsequences of the genetic abnormalities underlying male subfertility (or infertility) and might become stimulators for the development of novel approaches and applications in the treatment of infertility. In addition, risks with a polygenic background appearing at birth as congenital anomalies and other theoretical or stochastic risks are discussed. Recent data suggest that assisted reproductive technology might also affect epigenetic characteristics of the male gamete, the female gamete, or might have an impact on early embryogenesis. It might be also associated with an increased risk for genomic imprinting abnormalities. (Asian J Androl 2006 Nov; 8: 643_673)
Keywords: genetic risks; epigenetic risks; intracytoplasmic sperm injection; testis; male infertility Correspondence to: Prof. Nikolaos Sofikitis, Department of Urology, Tottori University School of Medicine, 36 Nishimachi, Yonago 683, Japan. Tel: +30-6944-3634-28, Fax: +30-2651-0970-69 DOI: 10.1111/j.1745-7262.2006.00231.x Contents
1 The importance of the evaluation of microscopic and macroscopic consequences of ICSI techniques 2 Strong evidence proves a genetic basis of several spermatogenic defects 3 Genetics of male infertility 3.1 Single gene disorders 3.1.1 Congenital bilateral absence of vas deferens due to CFTR mutations 3.1.2 Kartagener syndrome and monomorphic abnormalities of spermatozoa 3.1.3 Genetic disorders with endocrine or neurologic implications 3.2 Chromosomal abnormalities 3.2.1 Autosomal translocations 3.2.2 Robertsonian translocations 3.2.3 Klinefelter syndrome 3.2.4 47,XYY syndrome 3.2.5 Structural abnormalities of the X-chromosome 3.2.6 Chromosomal inversions 3.3 Deletions of the Y chromosome 3.4 Evaluating chromosomal abnormalities in the gametes of males participating in ICSI programs 3.5 Mitochondrial aberrations of spermatozoa and ICSI 3.6 Reported congenital abnormalities and neuro-phychiatric development in children born after ICSI 3.7 Risks for chromosomal abnormalities in ICSI children 3.8 Exogenous DNA and HIV transmission risks from employment of ICSI procedures 3.9 Genetic and epigenetic risks from intraooplasmic injections of in vivo produced spermatids 3.10 Genetic risks after assisted reproduction techniques using in vitro generated male haploid germ cells 3.11 Epigenetic risks related to assisted reproduction techniques 3.12 Risks concerning transgenerational transmission of an acquired genetic or epigenetic defect 3.13 Risks related to mutations of genes regulating the spermiogenesis process 3.14 Preimplantation Genetic Diagnosis (PGD)-Biopsy techniques and Risks 4 Guidelines and Conclusions 1 The importance of evaluation of microscopic and macroscopic consequences of intracytoplasmic sperm injection (ICSI) techniques
ICSI represents a revolutionary technique of in vitro fertilization (IVF) developed during the past decade. It might represent the laboratory method of choice for the treatment of severe cases of male infertility. This method has become popular through the years and has been an invaluable stimulator for the development of novel approaches and applications along with the standard IVF. The use of ICSI resulted in the application of sympromatic (i.e. non-etiological) modes of treatment of severe cases of male infertility. In addition, ICSI has been a successful procedure for the fertilization of in vitro matured human oocytes [1]. Nevertheless, reservations for the effect of ICSI on the genetic constitution of the offspring derived from this technology have been raised [2]. Until the introduction of ICSI procedures in human assisted reproduction, the lack of an adequate number of competent spermatozoa for the performance of assisted reproduction methods (i.e. IVF) was a barrier for the achievement of pregnancies in cases where genetic deficiencies affected the male reproductive potential. However, nowadays, because ICSI techniques bypass several barriers in the natural fertilization process, there is much concern on the safety of ICSI and the probable transmission of reproductive deficiencies (of genetic etiology) or other genetic abnormalities to the offspring. Furthermore, the rapid employment of these methods in humans and the lack of organized experimental and clinical trials prior to the wide application of ICSI procedures have raised some additional concerns. One negative consequence of the use of ICSI techniques is the shift away from research on micro-insemination systems. Thus, there might be a need to develop new research directions. One new target might be the development of more stringent spermatozoal selection/preparation methods to reduce the risk of transmission of male genetic factors that have been associated with genetic risks for the ICSI offspring to the female gamete. In order to appreciate the potential genetic risks of ICSI techniques, it is necessary to analyze the causes of male infertility, particularly those that have a genetic basis. The use of ICSI procedures for the therapeutic management of infertile males with a genetic defect might overrun the limitations for transfer of this particular defect to the next generation. Thus, ICSI techniques might be responsible for the transmission of a genetic defect to the next generation. Therefore, ICSI procedures might propagate (i.e. maintain and increase) the incidence of a genetic defect related to the development of impaired spermatogenesis within a male population. Furthermore, because gametes and early embryonic genomes undergo an epigenetic reprogramming, ICSI techniques might interfere with the establishment of normal parental imprinting, resulting in embryonic or fetal abnormalities [3, 4].
2 Strong evidence proves a genetic basis of several spermatogenic defects During the past decade, there has been a dramatic expansion in the number of genes involved in spermatogenesis, sexual differentiation and reproductive deficiencies. The development of differential display reverse transcriptase-polymerase chain reaction (RT-PCR) procedures has led to the identification of many genes that are differentially regulated in various cell and tissue types [5]. Anway et al. [5] used the above technique to identify genes that are expressed in isolated mouse testicular type A spermatogonia and in more advanced germ cells. The authors identified cDNA fragments for mDEAH9, RanBP5, GC3, GC12, and GC14 genes in the testis and type A spermatogonia from wild type mice but not in samples from mutant sterile W/Wv mouse testis. RT-PCR analyses of isolated spermatogonia, pachytene spermatocytes and round spermatids found that mDEAH9, RanBP5, GC3, GC12 and GC14 genes were expressed in all three cellular populations. RanBP5 expression appeared to be regulated during the cycle of the seminiferous epithelium with the highest expression in stages III through VII. Expression of GC14 was greatest in the meiotic germ cellular subpopulations. In addition, Anway et al. [6] identified a murine testis complementary DNA encoding a homolog to human A kinase anchoring protein-associated sperm protein (ASP). Northern blot and RT-PCR analyses did not detect ASP mRNA in mouse spleen, brain, liver, lung, heart, kidney, skeletal muscle, ovary or Sertoli cells. In contrast, the above techniques localized ASP mRNA to the germ cell compartment of the seminiferous tubules in the testis. In addition, Anway et al. [7] provided strong evidence that the effects of endocrine disruptors on spermatogenetic capacity in subsequent (F1 and F2) generations might be the result of altered DNA methylation patterns in the male germ line. The latter study showed the ability of environmental factors to reprogram the genes in the male germ line and to promote a transgenerational disease state [7]. Other studies by Anway and Skinner [8] confirmed the transfer of abnormal phenotypes (through epigenetic actions on the male germ line) to subsequent generations analyzed. Mouse models with reproductive defects as a major phenotype have been created and now hold over 200 [9]. These models are helping to define mechanisms of reproductive function, as well as identify potential new genes involved in the pathophysiology of reproductive disorders. Mouse models for the study of reproductive defects have been produced by spontaneous mutations, transgene integrations, retroviral infection of embryonic stem cells, ethylnitrosurea mutagenesis and gene targeting technology. Several genes required for vertebrate fertility are highly conserved in evolution with orthologues in Drosophila melanogaster (i.e. DDX4), fat facets (DFFRY), and boule (DAZ) [10_12]. Defects in sexual differentiation pathways can cause infertility in mice and humans of both sexes. It has been pointed out by Matzuk and Lamb [9] that several gene defects or gene-related pathophysiologies leading to defects in sex determination or development (i.e. pseudohermatidism, sex reversal, Denys-Drash syndrome, pseudovaginal perineoscrotal hypospadias, cryptorchidism or congenital bilateral absence of vas deferens), defects in sperm production and function (i.e. myotonic dystrophy, Nooman syndrome, sickle cell anemia, b-thalassemia, Kartagener syndrome, primary ciliary dyskinesia, Fanconi anemia or ataxia telangiectasia) and endocrinopathies lead to human male infertility. In addition, numerical/structural chromosomal abnormalities result in human male infertility as well. Knockout animal models have provided strong evidence supporting the genetic basis of human male infertility in subpopulations of infertile men. Of major importance are research efforts focused on the genes of sex chromosome Y and also on genes associated with certain genetic syndromes having the development of male infertility as an inherent component of their phenotype. Consequently, these studies provide evidence for the molecular basis of the genetic risks of ICSI procedures. Today, a significant percentage of spermatogenic abnormalities can be studied and classified according to genetic criteria. In fact, 30% of spermatogenic abnormalities are considered to have a genetic basis-related etiology [13_15]. A clinical classification of spermatogenic disorders alone cannot directly associate a phenotype with a particular genetic abnormality. Excluding the genetic syndromes/pathophysiologies showing infertility as one of the characteristics of their clinical phenotype, in the vast majority of infertile males the clinical diagnosis of infertility is not associated with any other clinically important phenotypic manifestations/characteristics. In most infertile males, the aetiology of infertility is unknown (i.e. idiopathic). This is the reason the majori-ty of fertility specialists recommend the clinical and laboratory evaluation of infertile males before the application of ICSI techniques. A major objective of the current communication was to associate the genetic defects of infertile males with their semen quality and reproductive potential. Another objective was to emphasize the probability of the transmission of major or minor paternal genetic defects to the embryo/offspring when ICSI procedures are applied. Major genetic or epigenetic defects in the male XY-embryo might be manifested at the fetal or neonatal stage as profound and severe manifestations [9, 16]. In contrast, minor genetic defects in the male XY-embryo might not affect the early embryonic development directly but might play a significant detrimental role in the reproductive potential of the affected newborns.
3 Genetics of male infertility
3.1 Single gene disorders A subpopulation of patients that present to IVF cli-nics for treatment of male factor infertility might have incomplete penetrance of a single gene genetic disorder. Another population might show some clinical manifestations characterizing the disorder that is the cause for the development of infertility.
3.1.1 Congenital bilateral absence of vas deferens due to cystic fibrosis transmembrane conductance regulator gene mutations Most of the congenital bilateral absence of vas defe-rens (CBAVD) cases (60_90%) and some cases of unilateral absence of the vas deferens are to the result of mutations of the cystic fibrosis transmembrane conductance regulator (CFTR) gene. This gene is responsible for the underlying genetic defect in cystic fibrosis (CF), a genetic recessive disorder with an incidence of carriers between 5_6% in the Caucasian population. Among infertile patients with CBAVD, the incidence of CFTR mutation-carriers is estimated to be 20-fold greater than that in the general population [17]. Mutations in CFTR are classified as severe or mild. The association between the genotype and the phenotype is complex. In general, the mild mutations result in mild alterations in phenotypes restricted in the male reproductive tract and are characterized by obstructive azoospermia. More than 700 mutations in CFTR gene spanning (approximately 230 kb) have been described [18]. CBAVD patients have either two mild CFTR mutations or a mild mutation in combination with a severe one. The most frequent severe mutation is the DF508 representing the majority (60_70%) of the CF mutations in carriers and patients. In addition, polymorphisms reducing the production of the CFTR protein (5T, 7T) have been shown. In particular, the homozygous or heterozygous presence of the 5T allele is a frequent finding in CBAVD patients with incomplete penetrance. The identification of this allele, corresponding to an inefficient acceptor splice site with a 90% reduction of the CFTR protein synthesized, is associated with a spectrum of presentations of phenotype from healthy fertile males to CBAVD patients [19]. Compound heterozygotes carrying the 5T allele but showing a CFTR mutation might present with atypical or typical clinical phenotypes of CF. At least seven other mutations commonly related to CBAVD have been described and they are almost all related to defective CFTR protein processing [17]. In addition, the missence R117H mutation in exon 4 is also related to CBAVD in association with the 5T variant [20]). Thus, testing for R117H and 5T/7T/9T polymorphism is important in the infertility setting. Recovery of epididymal or testicular spermatozoa and subsequent employment of ICSI techniques are essential to assist reproduction in the group of CBAVD male patients. This approach has the risk of producing affected offspring when the female partner is a carrier. Consequently, at least the most common CFTR mutations (up to 90%) should be screened (see above paragraph). Genetic counselling is strongly recommended for these patients (Table 1). Testing the obstructed azoospermic men for the most common mutations and associated polymorphisms (28 in total) is the appropriate first step. Preimplantation genetic diagnosis (PGD) is recommended for couples who are both positive for CF mutations and wish to integrate ICSI and genetic diagnosis at early stages of the embryonic development [21, 22]. Josserand et al. [23] detected CFTR mutations on 56 alleles of 50 males with congenital bilateral absence of vas deferens. A total of 15 (30%) were compound heterozygote and 26 (52%) heterozygote. In all, 38% of the patients had a positive sweat test. It appears that congenital absence of vas deferens can be seen in male heterozygote carriers of one CFTR mutation or compound heterozygotes with two mutations, one of which might not be detected by the mutation analysis. This is important, as it will affect counselling of couples especially if the female partner carries a CFTR mutation. 3.1.2 Kartagener syndrome and other monomorphic anomalies of spermatozoa Primary akinesia or dyskinesia of the cilia is a gene-ral term used to describe disorders of the structure of the cilia mainly in the airways and the sperm tail resulting in impaired sperm motility [24]. Affected individuals have chronic manifestations (as a result of the above disorder) in their airways. Males are usually infertile as a result of the sperm tail defects. There are structural anomalies in the proteins forming the bridging links of the dynein in the axoneme [25]. The co-existence of sinusitis, bronchiectasia, immotile spermatozoa and situs inversus characterizes Kartagener syndrome. The prevalence of situs inversus of any etiology appears to be in a range between 1 in 25 000 and 1 in 8 000. Twenty to 25% of these individuals with complete mirror-image situs inversus have ciliary dyskinesia and respiratory symptoms (Kartagener syndrome) as associated findings [26]. The prevalence of Kartagener syndrome in the general population is approximately 1: 40 000. Earlier linkage analyses in a large number of primary ciliary dyskinesia families showed extensive heteroge-neity [26]. No single genomic region harbouring a common primary ciliary dyskinesia locus was identified. However, several potential chromosomal regions that could harbour genes for primary ciliary dyskinesia were localized [26]. To date, mutations in two genes have been associated with a minority of primary ciliary dyskinesia/Kartagener syndrome cases. These are genes coding for the dynein axonemal heavy chain 5 and the dynein axonemal intermediate chain 1. A considerable number of additional monomorphic human sperm defects have been described. Most appear to be exceedingly rare and they might only be detectable through electron microscopy [27]. For the `9 + 0' axoneme defect [28] and globozoospermia (round head defect), evidence from family studies suggests that these are genetically determined disorders [29]. The mode of inheritance of monomorphic human sperm defects is most likely to be autosomal recessive or X-linked [13]. No mapping data for the responsible genes are available yet [13]. Thus, monomorphic anomalies of spermatozoa represent a defined entity with distinct genetic background and variable characteristics as, for example, globozoospermia [13, 24] (see the section 3.13). Globozoospermia is found in less than 0.1% of infertile male partners [30]. Although these pathophysiologies of sperm motility and morphology are heterogenous, the genetic diagnosis is based on the clinical and laboratory examination, and the appropriate genetic tests (see the section 3.13). In a recent study, no mutation was found among six patients with globozoospermia [30]. Counseling is of paramount importance to inform the couples about the risk of transmitting these disorders to their offspring.
3.1.3 Genetic disorders with endocrine or neurologic implications Kallman syndrome is implicated in approximately 5% of the infertile males with hypogonadotrophic hypogo-nadism. Anosmia is a result of deletions in the Xp22 region or mutations of the KAL-1 gene. The syndrome phenotype varies from normogonadotrophic fertile patients to the total absence of the gonadotrophins (FSH and LH) as a result of insufficiency of GnRH. The full abnormal phenotype is due to the inefficient migration of the hypothalamic olfactory neurons and those producing GnRH. When the serum testosterone profiles are sufficient to support sexual differentiaton, the male phenotype is normal and spermatogenesis can be stimulated by gonadotrophins to permit subsequent use of ICSI procedures [31]. GnRH receptor gene mutations (autosomal recessive inheritance) result in hypogonadotropic hypogonadism with oligospermia. In addition, FSH receptor gene mutations are associated with variable degrees of spermatogenic defects. Activating mutations of the same gene have been described. Furthermore, mutations in genes encoding the LH receptor, 5a-reductase 2, or CYP 21 might cause defects in spermatogenesis [32]. Affected males might be treated with ICSI and, therefore, are at risk to transmit the underlying defect to the offspring. A form of Kennedy disease characterized by androgen resistance and a molecular defect in the androgen receptor gene is associated with male infertility and defects in spermatogenesis [33_35]. The main feature of this condition is spinobulbar muscular atrophy (SBMA) with neurodegenerative phenotype. The gene responsible for the expression of androgen receptor is located on the X chromosome (Xq11-q12, OMIM #313700). The latter men might be candidates for ICSI techniques before the full onset of their disease, and they should also be informed that the consequences of their disease might be considered much more devastating than the infertile phenotype and that their disease might result in severe clinical manifestations. Nevertheless, as we have previously reported, couples with female SBMA carriers might request PGD in order to assure the birth of an unaffected offspring [36]. Myotonic dystrophy and fragile X syndrome, similarly as the Kennedy disease, represent disorders characterized by dynamic trinucleotide repeat expansions. Decreased sperm function or azoospermia are common in patients with myotonic dystrophy [37_39]. In cases of myotonic dystrophy of intermediate clinical severity, the use of combined ICSI and PGD procedures might assist to prevent the transmission of the defect to the offspring [40]. The X chromosome is not transmitted directly through a male carrier of an X-linked disorder to his male offspring, nevertheless it can be transmitted via a daughter to a male grandchild. Sermon et al. [40] have described their experience with fluorescent PCR and automatic fragment analysis for the clinical application of pre-implantation genetic diagnosis of myotonic dystrophy. The prevalence of the fragile X syndrome (FRAXA) premutation carriers is 1/1 000 in males and 1/350 in females, whereas the prevalence of full mutation is 1/4 000 males or females [41]. Carriers of premutations have mild or no symptoms, whereas male patients with full mutation of the FRAXA syndrome have moderate to severe mental retardation, behavioural problems and spermatogenic impairment including abnormal tubular morphology and excessive number of malformed spermatids. The overall result is decreased fertility probably as a result of the fact that the gene that is responsible for the phenotype is expressed in the male gonads [42, 43]. The use of ICSI procedures as a treatment for males with FRAXA syndrome mutations, or even permutations, is definitely susceptible to serious ethical considerations. Couple counseling, written consent forms and, probably, National Authority Permission is necessary. Platteau et al. [44] claimed that PGD work-up for FRAXA syndrome couples should include a determination of the premutation or mutation carrier status and the paternal or maternal origin of the premutation/mutation. Fragile X-premutation carriers should be advised not to postpone reproduction. Female premutation carriers have up to 50% (depen-ding on CGG repeat size) risk of fragile X syndrome in their offspring and a risk (15_20%) of premature ovarian failure [41, 45]. Up to 30% of females with a full mutation can be symptomatic depending on the X-inactivation status. Female premutation carriers belonging to families with fragile X syndrome should ask for PGD or prenatal diagnosis (PND) in order to prevent transmission of the disease [46]. Sermon et al. [46] reported for the first time in the literature a method for PGD for FRAXA syndrome based on the amplification of the CGG triplet in the normal allele. The above-mentioned single gene genetic disorders indicate the risks of transmitting genetic abnormalities via ICSI procedures and stress the need for systematic genetic testing in familial or sporadic infertility cases (Table 1).
3.2 Chromosomal abnormalities Chromosomal abnormalities have been associated with infertility or subfertility in males. The incidence of chromosomal abnormalities in the karyotypes of infertile males is 5.8%, with a predominance of sex chromosomal abnormalities according to a review of pooled data from 11 surveys (9 766 men with azoospermia or oligospermia were evaluated) [2, 47]. The phenotypic consequences of the sex chromosomal abnormalities are usually mild compared with the consequences of autosomal chromosomal abnormalities in males [14]. In addition, the incidence of chromosomal aneuploidies, especially those shown in the sex chromosomes, is higher in spermatozoa from men with non-obstructive azoospermia [48]. Mateizel et al. [49] have shown that aneuploidy for chromosome 18 is more frequent in men with spermatogenic failure. Furthermore, sperm concentrations smaller than 20 × 106 spermatozoa/mL are associated with significantly higher percentage of de novo chromosomal anomalies in prenatal samples in successful pregnancies [50, 51]. Numerical abnormalities of the sex chromosomes might be found either in immature testicular germ cells (germline defects) or in spermatozoa of men whose peripheral blood cytogenetics indicate non-mosaic Klinefelter syndrome (gonadal mosaicism) [52]. If ICSI procedures are scheduled for the therapeutic management of male infertility associated with chromosomal abnormalities of the male partner, it is important to discuss with the couple the option of PGD or PND (Tables 1, 2).
3.2.1 Autosomal translocations Autosomal translocations are 4_10 times more frequent in infertile (subfertile) males compared with fertile individuals [53, 54]. Mendelian Cytogenetic Network has approximately 265 entries of balanced reciprocal tranlocations from infertile males [55]. Among balanced chromosomal rearrangements in male infertility, half of the identified autosomal breakpoints (5/10) were found to be located on chromosome 1, suggesting a clustering of male specific loci on this chromosome. The above breakpoints along chromosome 1 have been found to be in excess in infertile males (from the Mendelian Cytogenetics Network) compared with the karyotypes of a cohort [56]. In general, reciprocal or non-reciprocal autosomal chromosomal translocations and complex chromosomal rearrangements (involving three or more chromosomes) are associated with subfertility. This is the result of inappropriate pairing of the homologous chromosomes during meiosis, leading to meiotic disturbance or chromosomal imbalance in the male gametes [2, 57, 58].
3.2.2 Robertsonian translocations Translocations between acrocentric chromosomes (Robertsonian) are frequent in humans, but their impact on spermatogenesis varies from the absence of spermatogonia to the development of normal spermatogenesis. The therapeutic management of Robertsonian translocations associated with infertility depends on the presence of spermatozoa and the success of ICSI procedures. In these cases, ICSI procedures raise risks for chromosomal abnormalities in the generated embryos [21, 22, 59]. The reproductive risks for the newborn, as a result of the presence of Robertsonian translocations in the infertile couple, depend on the chromosomes involved and the sex of the carrier. The most common risks are related to newborn translocation trisomies of chromosomes 13, 14, 21 or 22. An increased proportion of carriers of robertsonian translocations (usually t[13q;14q]) has been reported among oligozoospermic (1.6%) and azoospermic (0.09%) men attending infertility clinics or among the male partners in couples with recurrent spontaneous abortions [2, 60]. Therefore, there is a strong indication for the performance of PGD in combination with the ICSI procedures [61]. For the evaluation of the chromosomal composition of spermatozoa, fluorescent in situ hybri-dization (FISH) techniques are recommended with additional (to the probes for sex chromosomes) specific probes for chromosomes participating in probable reciprocal or Robertsonian translocations [62_64]. Van Assche et al. [63] carried out PGD and sperm analysis by FISH for the most common reciprocal translocation t (11:22). By choosing probes lying on both sides of the breakpoints and by using a combination of subtelomeric or locus-specific probes and centromeric probes, the use of three-color FISH enabled detection of all the imbalances in sperm and/or cleavage stage embryos in the patients.
3.2.3 Klinefelter syndrome Non-mosaic Klinefelter (47,XXY) and mosaic Klinefelter syndrome (46,XY/47,XXY) are the most common chromosomal abnormalities observed in azoospermic males. Adult males with non-mosaic Klinefelter syndrome (47,XXY) have hypogonadism and infertility. Disruption (arrest) in spermatogenesis is shown. Spermatogonia in these patients usually do not further differentiate beyond the stage of primary spermatocyte, but occasionally testicular focal advanced spermatogenesis up to the spermatozoon stage is observed. FISH analysis of spermatogonia and spermatocytes from men with non-mosaic Klinefelter syndrome show a variable frequency of aneuploidy of the sex chromosomes (either 47,XXY or 46,XY profiles are shown indicating gonadal mosaicism) [52, 65, 66]. Spermatozoa recovered from testicular biopsies of men with karyotypes indicating non-mosaic Klinefelter syndrome have been used to fertilize oocytes by ICSI techniques. Preimplantation blastomere-FISH analysis should be carried out with X and Y probes to confirm that the sex chromosomal complement of the embryos that are going to be transferred is normal. The birth of normal offspring has been reported after ICSI techniques using testicular spermatozoa recovered from men with non-mosaic Klinefelter syndrome [52, 65, 67-69; among others]. We can speculate that the risk of transmitting additional X chromosomes to the offspring might be related to the percentage of the 24,XY testicular spermatozoa in the recovered testicular sperm population. It appears logical to speculate that a man with a non-mosaic Klinefelter syndrome and a large percentage of abnormal 24,XY spermatozoa in his testicular biopsy sample he may have a large probability to generate a 47,XXY embryo after ICSI techniques. A number larger than 20 human offspring have been fathered by men with non-mosaic Klinefelter syndrome [52, 65]. Although all the latter offspring are normal (46,XY or 46,XX), PGD or PND are strongly recommended. Ron-El et al. [70] have reduced a 47,XXY embryo implanted after ICSI and embryo transfer techniques in a couple with Klinefelter syndrome. Previous studies in our laboratory have shown that among men with non-mosaic Klinefelter syndrome, those with larger secretory function of Sertoli cells have a higher probability to be positive for testicular foci for spermatogenesis up to the spermatozoon stage [52, 65]. In addition, we have previously shown that within a population of men with non-moaic Klinefelter syndrome, the larger the testicular telomerase profiles are the higher the probability of finding testicular spermatozoa is [52, 65]. In a recent study, Akashi et al. [71] reported a male patient with mosaic Klinefelter syndrome whose ejaculated spermatozoa were identified as being haploid by FISH before ICSI leading to the successful pregnancy of his wife and the birth of a healthy baby girl. When semen samples in men with either mosaic or non-mosaic Klinefelter syndrome are negative for spermatozoa, testicular biopsy should be carried out to recover haploid male gametes [52]. Although testicular fine needle aspiration has been used as a diagnostic tool in a general group of non-obstructed azoospermic men [72], its role in men with Klinefelter syndrome has not been evaluated. A subpopulation of men with non-mosaic Klinefelter syndrome has both 46,XY spermatogonia/primary spermatocytes and 47,XXY spermatogonia/primary spermatocytes in their seminiferous tubuli [52]. A previous study in our laboratory has not indicated sex chromosomal non-disjunctions during the meiotic divisions of the 46,XY spermatogonia/primary spermatocytes in men with non-mosaic Klinefelter syndrome [52]. Subsequently, similar numbers of testicular 23,X round spermatids and 23,Y round spermatids are thought to have been produced from the meiosis of the normal 46,XY spermatogonia/primary spermatocytes in the above men. To explain the larger proportion of 23,X round spermatids compared with the 23,Y round spermatids within a population of men with non-mosaic Klinefelter syndrome, an attractive speculation is that an XX pairing and a univalent Y chromosome type of pairing occurs in the great majority of 47,XXY primary spermatocytes that undergo regular meiosis [52]. In contrast, an XY pairing and a univalent X chromosome type of pairing might occur in a minority of 47,XXY primary spermatocytes that undergo regular meiosis. This speculation can explain a) the increased proportion of the hyperhaploid 24,XY round spermatids compared with the hyperhaploid 24,XX round spermatids within a population of men with non-mosaic Klinefelter syndrome [52], and b) the larger proportion of testicular 23,X round spermatids compared with testicular 23,Y round spermatids within a population of men with Klinefelter syndrome [52, 65]. XX pairing and a univalent Y type of pairing in 47,XXY primary spermatocytes that undergo meiosis is expected to result in increased proportions of 23,X round spermatids/spermatozoa and 24,XY round spermatids/spermatozoa (post-meiosis) in the testicles of men with Klinefelter syndrome [73]. This is because a regular meiosis in a 47,XXY spermatogonium with an XX pairing and a univalent Y should lead to the production (from one 47,XXY spermatogonium) of two 23,X spermatids and two 24,XY spermatids [73]. Increased proportions of 24,XY round spermatids compared with 24,XX round spermatids within a population of men with Klinefelter syndrome and larger proportion of 23,X round spermatids compared with 23,Y round spermatids have been found, indeed, within a population of men with non-mosaic Klinefelter syndrome in our laboratory [52]. In contrast, if a XY pairing and a univalent X had been present in the majority of 47,XXY primary spermatocytes, regular segregation of the sex chromosomes would have resulted in increased proportions of a) 23,Y round spermatids/spermatozoa (compared with 23,X round spermatids/spermatozoa) and b) 24,XX round spermatids/spermatozoa (compared with 24,XY round spermatids/spermatozoa) in the testicles of men with Klinefelter syndrome [73]. In fact, if a XY sex vesicle is formed and the extra X chromosome is free, regular segregation of the sex chromosomes would produce (from one 47,XXY primary seprmatocyte) two 24,XX spermatids/spermatozoa and two 23,Y spermatids/spermatozoa [73]. It appears that the findings of our previous study demonstrating an increased proportion of 24,XY round spematids compared with 24,XX round spermatids and a larger proportion of 23,X round spermatids compared with 23,Y round spermatids suggest an XX pairing a Y univalent in the majority or in all of the 47,XXY primary spermatocytes that undergo meiosis [52]. Therefore, we might suggest that an XX pairing and a univalent Y chromosome type of pairing occurs in the great majority of 47,XXY primary spermatocytes that undergo meiosis.
3.2.4 47,XYY Paternal non-disjunction of the sex chromosomes during meiosis is the underlying cause for the presence of an extra Y chromosome. Although some 47,XYY males are fertile and produce normal gametes, a limited subpopulation of 47,XYY males might have severely impaired sperm production [74]. Although the additional Y chromosome might be spontaneously corrected during meiosis, there is a high incidence of disomic spermatozoa with 24,XY or 24,YY constitution [75]. Post-fertilization, the risk of aneuploidy of the sex chromosomes in the derived embryos might be expected to depend on the frequency of the aneuploid spermatozoa in the testicular tissue of the ICSI participants. It appears logical to speculate that the larger the percentage of sperm aneuploidies is within a population of testicular spermatozoa recovered from a testicular biopsy sample of a man with 47,XYY syndrome syndrome, the larger the probability is that the embryologist will aspirate and process for ICSI an aneuploid spermatozoon, with an overall result a larger probability to generate an aneuploid embryo. ICSI procedures are applicable with the reservation of a higher genetic risk for aneuploid embryos. PGD or PND are strongly recommended.
3.2.5 Structural abnormalities of the X chromosome Structural abnormalities of the X chromosome, such as minor deletions or reciprocal translocations involving the chromosome X and an autosomal chromosome, are occasionally the cause of male infertility [76]. Deletions of a large part of the X chromosome of the female gamete results in the loss of one or more genes and is incompatible with the development of a male embryo after ICSI procedures because males have only one X chromosome and the loss of any genes normally located on the X chromosome is not compensated [14]. The results of an X-autosome translocation vary considerably depending on the sex of the carrier of such an aberration and the position of the translocation break points. Female carriers of a balanced X-autosome translocation generally are phenotypically normal. An important exception is evident in those women in whom the break points in the X chromosome involve the critical region Xq13-q26. These women are always infertile because of gonadal dysgenesis [77]. Reciprocal X-autosome translocations affect male fertility. A possible hypothesis is that reciprocal X-autosome translocations might interfere with X chromosome inactivation [77, 78]. Thus, it has been proposed that X-autosome translocations interfere with the process of X chromosome inactivation resulting in meiotic arrest at the primary spermatocyte stage. A probable hypothesis is the reactivation of the X chromosome, which is supposed to remain transcriptionally silent during spermatogenesis and the overall result, might be azoospermia [79, 80]. Information on the percentage of male germ cells with X-autosomal translocations in the above men is not available in the literature today. ICSI procedures might be applied in these cases (using testicular spermatozoa from testicular foci of advanced spermatogenesis) [14], however, there is a risk of transmission of either balanced or unbalanced chromosomal translocations in the resulting embryos. Production of secondary spermatocytes and spermatids (Figure 1) depends on the X chromosome inactivation driven by an X-linked gene acting at the primary spermatocyte stage. The X and Y chromosome form a single mass in the zygotene stage during pairing of the chromosomes at meiosis I [78, 81]. The pyruvate dehy-drogensa 1 gene is silent in spermatocytes and spermatids [80]. The inactivation of the X chromosome is essential to prevent the recombination between X and Y chromosomes during meiosis [80]. It is not clear why the X-chromosome should be inactivated during spermatogenesis. Because there is no evidence that pro-ducts of the X-chromosome are not permissive for spermatogenesis, it might be suggested that inactivation of the X-chromosome might reflect not the metabolic needs of the testicular germ cells but specific meiotic events such as chromosomal pairing and recombination. X-chromosome inactivation might be directed by an X-linked gene during the primary spermatocyte stage [14]. Thus, the existence of translocations involving the chromosome X might have a considerable effect in spermato-genesis, impairing the capacity of primary spermatocytes to enter meiosis [80]. In some cases, spermatogenesis progresses to the stage of elongated spermatids but this process is extremely inefficient and only a small number of spermatozoa is produced [14]. In patients having spermatids or few spermatozoa in testicular biopsies, the probability of chromosomal abnormalities in the embryos derived by ICSI techniques cannot be excluded. PGD might help to avoid transfer of the affected embryos [21, 22].
3.2.6 Chromosomal Inversions Inversions (peri- and paracentric) of chromosomes 1, 3, 5, 6, 9, 10 and 21 have been described in infertile men [60, 82_84]. The impact of chromosomal inversions in the development of impairment in spermatogenesis in infertile males is variable. Arrest at the primary spermatocyte stage has been described for a particular pericentric inversion on chromosome 1, whereas pericentric inversions of other chromosomes have been associated with azoospermia or oligospermia [60, 82]. The couples should be informed about the probability of spontaneous abortion if pregnancy is achieved via assisted reproduction [85].
3.3 Deletions of the Y chromosome Abnormalities in the Y chromosome are discussed separately in the present review study because the structural abnormalities of this chromosome have a direct effect on sexual differentiation and fertility. Various structural abnormalities of the Y chromosome are distinguishable at the molecular or the cytogenetic level. Translocations and microdeletions are the most frequently observed structural abnormalities. The Y chromosome is a complex chromosome that contains heterochromatin located among repeated genes, gene families and palindromic motifs. The non-recombining region of the Y chromosome contains three classes of euchromatic sequences [86], including: i) those that are transposed from the X chromosome during the process of the evolution of the Y chromosome (X transposed); ii) those sequences that are somewhat similar to sequence information from the X (X degenerate); and iii) those sequences that are repeated across the proximal short arm of the Yp and across most of the Yq. Translocations between the Y chromosome and autosomal chromosomes [87_89] appear to be more common and have a detrimental influence on spermatogenesis. Ooplasmic injections have been applied in such cases after testicular biopsy and recovery of spermatozoa. A risk of developmental delay as the result of chromosomal imbalance in the offspring has been suggested [90]. It has also been suggested (by a limited number of studies) that dicentric Y chromosomes do not allow spermatogenesis to proceed further than primary spermatocyte stage (early maturation arrest) [91, 92]. Therefore, ICSI procedures cannot be taken into consideration for the therapeutic management of these couples. In the Yq11.21-23 region, where the azoospermia factor (AZF) is located, there are three loci related to spermatogenesis (AZFa, AZFb and AZFc). These loci have been clustered in tandem and contain putative or candidate genes detrimentally affecting spermatogenesis when they are absent. In a general population of ICSI participants, the frequency of deletions is 2_3%, whereas in infertile males with azoospermia, the frequency of deletions is 6_12% [15, 93]. Deletions are present in 5.8% of men with severe oligozoospermia. Katagiri et al. [86] have shown an incidence of Y chromosome microdeletions equal to 16% in a population of azoospermic men and equal to 4% in a population of severe oligospermic men. In the above study, Y chromosome microdeletions were absent when sperm concentration was larger than 5 000 000 spermatozoa/mL. AZFa region harbors the genes DFFRY, USP9Y and DBY that are important for spermatogenesis. However, the most common deletions occur in AZFc and AZFb regions involving the DAZ and RBM multiple copy genes and other genes such as CDY1, PRY, TTY2 and EIF1AY expressed solely in the human testis [94, 95]. There is no clear association between the length of the deletion and the semen quality or the testicular histology. The phenotype varies from oligospermia to azoospermia with/or without testicular foci of spermatogenesis up to the spermatozoon stage. All patients with complete deletion of AZFa region or complete deletion of the AZFb region are azoospermic and negative for foci of testicular spermatozoa [96]. A strict genotype-phenotype correlation is observed only for the deletion of the entire AZFa and AZFb regions, which are associated with Sertoli cell-only syndrome and arrest at the primary spermatocyte stage, respectively [97]. On the contrary, the deletion of the most distal AZFc is associated with a heterogenous phenotype in different individuals ranging from the absence of germ cells in the testis to a severe reduction of the sperm number/motility/morphology in the ejaculate [98]. This phenomenon suggests that although spermatogenesis might start without AZFc genes, their presence is crucial to obtain quantitatively and qualitatively normal spermatogenesis. This region contains a total of eight gene families: BPY2, CDY1, DAZ, TTY3.1, TTY4.1, TTY17.1, CSPG4LY and GOLGA2LY. The classical AZFc deletion, which removes 3.5 Mb between the b2/b4 amplicons, is the most frequent type of deletion. A partial deletion termed gr/gr has been described in infertile men with varying degrees of spermatogenic failure. This deletion removes half of the AZFc region content. Another deletion with the name b2/b3 appears to have no effect on fertility status in association with a certain Y chromosome background commonly present in northern European populations [99]. The first multicopy gene identified in this region (i.e. AZFc) was the DAZ, which belongs to a gene family that consists of the two autosomal single copy genes BOULE and DAZL gene and the Y specific DAZ. No mutations for the DAZL and BOULE genes have been reported so far, except two single nucleotide polymorphisms in the DAZL gene [100]. Katagiri et al. [86] have reported surgical retrieval of epididymal spermatozoa from a man with partial deletion in AZFb region. His son had an identical deletion. Patients with AZFc deletions are either azoospermic (with or without testicular foci of spermatozoa) or have spermatozoa in the ejaculate. Additional studies confirmed that azoospermic men with complete deletions of either the AZFa or AZFb regions never demonstrated testicular spermatozoa after testicular biopsy procedures [101]. Testicular spermatozoa of men with (either complete or partial) AZFc deletions or partial AZFb deletions are anticipated to successfully fertilize oocytes and generate offspring at the same rate as non- deleted infertile men. In addition, a subpopulation of men with AZFc deletions has a certain degree of oligospermia that requires ICSI. The pathogenetic role of Y-chromosome deletions in male infertility has been questioned by reports describing microdeletions in proven fertile men [97]. However, male fertility is not a synonym for normozoospermia [97]. The pathogenetic significance of Y chromosome microdele-tions is spermatogenic failure and not infertility. In rare cases, transmission of an AZFc deletion has been reported via natural conception from a subfertile younger father to an infertile son [102]. Kuhnert et al. [103] reported natural transmission of an AZFc Y chromosome microdeletion from a father to his sons. Rolf et al. [104] have reported natural transmission of partial AZFb deletion over three generations. Kamische et al. [105] reported transmission of a Y-chromosomal deletion involving the DAZ and CDY1 genes from father to son through ICSI. Men with Y chromosomal microdeletions who are positive for spermatozoa will almost certainly pass the deletion to male offspring generated by ICSI procedures [106_109]. A progressive decrease in testicular spermatogenetic activity over time has been reported in some infertile men with AZFc microdeletions. Thus, testicular or ejaculated spermatozoa cryopreservation might be recommended for the latter men. Patsalis et al. [110] have suggested that there might be a potential risk of chromosomal aneuploidy for male offspring born to fathers with Y-chromosome microdeletions. This risk might include not only 45,X/46,XY offspring but also 45,X offspring. In addition, the above investigators recommended that PGD should be offered when men have ICSI for hypospermatogenesis caused by Y chromosome microdeletions to avoid transfer of 45X embryos. Data by Sofikitis et al. [111] using the testicular androgen-binding protein activity as a marker of Sertoli cell secretory function, does not show a defect in Sertoli cell secretory function in men with Y chromosome micro-deletions. We have previously hypothesized that in the future, it might be possible to achieve survival and differentiation of germ cells from non-obstructed azoospermic men (without genetically based causes of azoospermia) into the seminiferous tubuli of recipient human individuals (with AZFc microdeletions) who are negative for testicular spermatozoa [111]. The attractive hypothesis is that the recipient human Sertoli cells and the intratubular biochemical environment will support the donor human germ cells to differentiate. The above hypothesis is supported by studies in animals showing that the intratubular environment from infertile recipients can support the differentiation of donor germ cells from infertile subjects [111]. Some azoospermic couples who have considered using donor spermatozoa might be attracted by the idea of achieving pregnancy via sexual intercourse, even if the male partner ejaculates donor rather than his own spermatozoa into the reproductive tract of the female partner. Even in Sertoli cell-only syndrome testicular histo-logy (in sections stained by hematoxylin_eosin) from subpopulations of men with Y chromosome deletions, there is a probability that spermatids or spermatozoa can be identified in seminiferous tubules. It has been estimated that spermatozoa (either in the ejaculate or the testicular tissue) can be found in approximately 50% of azoospermic men with microdeletions in the AZFc region of the Y chromosome. Because AZF microdeletions are transmitted from the father to the male offspring, genetic evaluation for Y chromosomal deletions is recommended in non-obstructed azoospermic men or severely oligoasthenospermic individuals. In addition, large microdeletions of the tip of the Yq chromosome might cause chromosomal instabi-lity and might be responsible for chromosomal rearrangements or even Y chromosome loss. Issues, such as testicular mosaicism of Y chromosomal deletions, expansion of the Y chromosome deletions in the offspring, lower fertilization rates post-ICSI and familial basis of Y deletions represent the target of several investigations but the results are still inconclusive [112]. Because ICSI techniques are commonly used in patients with Y chromosome microdeletions, thus posing a considerable risk of passing the deletion on to the offspring [113], proper genetic counseling followed by detailed family history and specific molecular or cytogenetic assays are recommended.
3.4 Evaluating chromosomal abnormalities in the gametes of males participating in ICSI programs Males with severe oligospermia, obstructive azoospermia or non-obstructive azoospermia with testicular foci of spermatogenesis up to the spermatozoon stage represent the majority of candidates for ICSI. Several studies have been focused on the chromosomal constitution of spermatozoa of fertile and infertile men using FISH procedures [114, 115]. Although there is a remarkable variability in the methodology of these studies (i.e. regarding the number of FISH probes used or the selection of the patients), the findings of all these investigations indicate chromosomal abnormalities in the spermatozoa of ICSI participants (either oligospermic or azoospermic with testicular foci of spermatozoa). These abnormalities are mainly diploidy, autosomal disomy and nullisomy or aneuploidies of the sex chromosomes [114]. Spermatozoa recovered from non-obstructed azoospermic men (with testicular foci of advanced spermatogenesis) do have a higher incidence of chromosomal aneuploidy patterns among which sex chromosomal aneuploidy is the most common [48_50]. Mateizel et al. [49] have shown that the frequency of aneuploidy for chromosome 18 was higher in a group of azoospermic men with spermatogenic failure than in a group of azoospermic men with normal spermatogenesis. Huang et al. [116] reported an increase in the frequency of sex chromosomal abnormalities in testicular spermatozoa of non-obstructed azoospermic men. In another study, Viville et al. [117] showed that in obstructed azoospermic men (with or without CFTR mutation), there have not been significant differences in the chromosomal constitution of testicular spermatozoa compared with normal semen samples. In subpopulations of infertile men with primary testicular damage as a result of non-mosaic Klinefelter syndrome, there is a significant increase in the proportion of spermatids/spermatozoa with chromosomal aneuploidies. However, the majority of spermatids/spermatozoa (if they are present in testicular biopsy material) in the latter men have the normal haploid constitution of the chromosomes [52]. In a recent study, there was no significant difference in the incidence of aneuploid embryos between couples with obstructive azoospermia and couples with non-obstructive azoospermia [118]. Nevertheless, in both groups of the above study, the percentage of aneuploid embryos was relatively high (53_60%), indicating the potential risks of the employment of testicular spermatozoa for ICSI treatment. These patients would require a systematic monitoring of spontaneous abortions or implantation failures. In addition, the ICSI treatment should be coupled with PGD or PND for early identification of chromosomally abnormal embryos.
3.5 Mitochondrial aberrations of spermatozoa and ICSI The presence of mitochondrial abnormalities in spermatozoa has been proposed to be a cause of male infertility; mitochondrial abnormalities have been associated with asthenospermia [119]. Low sperm motility might be associated with deformations of the mitochondrial sheath containing functional mitochondria. The combination of fluorescence microscopy and flow cytometry with electron microscopic investigations is a sensitive, precise and comprehensive examination which helps discover sperm mitochondrial abnormalities that cause asthenozoospermia [119]. Successful ICSI in a case of severe asthenozoo-spermia that is the result of non-specific axonemal alterations and abnormal or absent mitochondrial sheaths has been reported [120]. The application of ICSI procedures in such patients implies introduction of the whole spermatozoon into the ooplasm and raises the question of potential risks for the derived embryo attributable to the transmission of paternally inherited abnormal mitochondrial DNA into the ooplasm of the oocyte. One study has evaluated the risk of heteroplasmy (mosaicism of paternal and maternal mitochondria) in 27 newborns born after ICSI procedures. Heteroplasmy was shown in a frequency of 0.1_1.5% (which is considered to be normal and so far does not appear to be alarming) [121].
3.6 Reported congenital abnormalities and neurophy-chiatric development in children born after ICSI Given the concerns from what has been already discussed in the present communication, it is important to analyze the outcome of some prominent ICSI programs and that of the ESHRE ICSI Task Force. The reported results from prenatal diagnoses in pregnancies achieved by ICSI techniques, indeed, showed a tendency for a higher frequency of aneuploidy of the sex chromosomes when compared with naturally conceived children [51, 67, 122_125]. Prospective data from Brussels have addressed the genetic consequences of the use of ICSI techniques in two consecutive studies evaluating 1 987 and 2 889 infants born after ICSI trials [51, 123, 126]. The outcome of ICSI techniques concerning the karyotypes, the existence of congenital abnormalities and the somatic or mental development was recorded. In total, 1.66% de novo chromosomal abnormalities of the autosomes and the sex chromosomes in equal proportions were found with an additional 0.92% of inherited structural chromosomal abnormalities (eight balanced and one inbalanced) from the father. Major congenital abnormalities were shown in a percentage equal to 2.3% of the total number of the children delivered. Fetal deaths were observed in a frequency of 1.1% after the 20th week of pregnancy. The second study compared the data between ICSI (n = 2 889) and IVF infants (n = 2 995) born in the periods 1991_1999 and 1983_1999, respectively. Using the same criteria and follow-up period, the ICSI group did not show an increased risk for major malformations or complications in comparison with the IVF group [51, 123]. Other studies comparing IVF with ICSI or ICSI-children versus children in a general population did not show any excess risk for ICSI children with the exception of the appearance of hypospadias (compared with the lower frequency of hypospadias in the general population), probably related to the paternal subfertility or to the hormones the mother received during the beginning of pregnancy [127, 128]. Although there is a subpopulation of non-obstructed azoospermic men where the etiology of azoospermia has a genetic basis [115, 129], there is no evidence for significantly higher risks for congenital abnormalities in infants born after ICSI procedures with epididymal or testicular spermatozoa (compared with naturally conceived offspring) [123,126,130_132]. Furthermore, replacement of frozen/thawed embryos generated by ICSI was not accompanied by a significantly higher incidence of congenital abnormalities in the newborns. In another report from Sweden, data concerning 1 139 children born after ICSI procedures were reviewed [127]. A consi-derable frequency of 7.6% of congenital abnormalities was observed and less than half of these abnormalities were minor. In that study, the relative risk of ICSI children to show a congenital abnormality was 1.75% but when this risk was corrected for twins or triplets it dropped to 1.19%. The only congenital abnormality with the alarmingly high relative risk of 3% was hypospadias. In other studies, the somatic development of children delivered post-ICSI techniques has been shown to be normal, whereas evaluation of mental development and fertility of the offspring need longer and more pervasive studies [125]. In order to reduce the potential risks of ICSI procedures for the fetus/newborn, cytogenetic analysis in haploid male gametes (recovered either from ejaculates or testicular biopsy samples) might be recommended before ICSI procedures are carried out in men with low sperm counts or in azoospermic men. Counseling and PGD or PND are of paramount importance. Mental and neuropsychiatric development in children delivered after ICSI techniques have been addressed in two successive reports. Both reports lacked a conclusion that supported a major abnormality in ICSI children or a significant deviation from the normally naturally conceived population apart from a) the findings concerning the presence of hypospadias [127, 128], or b) the complications related to multiple gestations [125, 130]. In a recent study [133], it was shown that singleton ICSI and IVF 5-year-olds are more likely to need health care resources than naturally conceived children. In addition, in that study, it was found that ICSI children presented with more major congenital malformations and both ICSI and IVF children were more likely to need health care resources than naturally conceived children. In another study [134], apart from a few interaction effects between mode of conception and and demographic variables, no differences were found when ICSI, IVF and naturally conceived scores on the WPPST-R and MSCA Motor Scale were compared. Nevertheless, the aforementioned interaction effects could indicate that demographic variables, such as maternal age at the time of birth and maternal educational level, play different roles in the cognitive development of IVF and ICSI children compared with naturally conceived children.
3.7 Risks and consequences of chromosomal abnormalities in ICSI children Pooled data from a survey of results of international trials point towards a slightly elevated frequency of sex chromosome abnormalities in ICSI children (compared to the general population). Overall ICSI results (in terms of percentages of chromosomal abnormalities in fetus karyotypes) do not appear to be significantly different compared with those of IVF [51, 123]. In general, the outcomes of IVF and ICSI trials are similar [51, 123]. The incidence of de novo numerical sex chromosomal anomalies in ICSI children ranges from 0.23_0.83%, which appears to be slightly higher compared with the 0.19% reported in the literature for the general population. De novo numerical autosomal chromosome abnormalities in ICSI children range from 0.5_1.4%. The latter percentage is 3 to 10 times higher than that in the general population (0.14%). Concerning the percentage of de novo structural chromosomal re-arrangements, there is a significant (3 to 4 times) increase from 0.07% in the general population to 0.23_0.27% in ICSI children [51, 123, 130_132]. In children born after ICSI techniques are carried out, most of these rearrangements are reciprocal and therefore do not have phenotypic consequences in the carriers. Never-theless, these rearrangements might be responsible for the generation of abnormal male gametes by meiotic malsegregation leading to chromosomally abnormal offspring postfertilization [130, 131, 135]. Male carriers of numerical or structural chromosomal abnormalities might father offspring with abnormal and meiotically incompetent cell lines at the age of reproduction after ICSI techniques [75, 136]. There are reports of low pregnancy rates in couples with primary testicular damage (after assisted reproductive technology), probably as a result of a generalized tendency of chromosomal nondisjunction [16]. In addition, ICSI with testicular spermatozoa has been proven to be less successful in men with non-obstructive azoospermia compared with men with obstructive azoospermia [137]. The increased chromosomal aneuploidy in testicular spermatozoa from men with non-obstructive azoospermia might explain the lower fertilization and pregnancy rates observed in that study [137]. Consistently, Aytoz et al. [138] have shown, after ICSI techniques, that within a group of couples that underwent ICSI techniques with ejaculated spermatozoa, the rate of intrauterine death was higher in a severely defective sperm subgroup than in better quality sperm subgroups. The higher percentage of chromosomal abnormalities in ICSI-children compared with the general population is probably related to the parental chromosomal abnormalities (mainly in the father) [51, 123, 125, 139]. This increase in chromosomal aberrations after ICSI procedures might also result from the selection of spermatozoa, which would otherwise be unable to naturally fertilize an oocyte [117, 126, 130_132]. In a study comprising a large number of prenatal tests carried out on pregnancies that were the result of ICSI techniques, a sixfold increase in sex chromosomal aberrations and a twofold increase in autosomal chromosomal aberrations was reported [130_132]. In additional studies, a significantly higher rate of de novo chromosomal abnormalities in amniocentesis was observed in ICSI offspring relating mainly to a higher number of sex chromosomal abnormalies and partly to a higher number of autosomal structural abnormalities [51, 123]. This finding was related to sperm concentration and motility of the ICSI participants. The significantly higher rate of observed inherited abnormalities in the ICSI prenatal tests compared with prenatal tests in the general population was related to a higher rate of constitutional chromosomal anomalies, mainly in the fathers [51, 123]. In addition, post-ICSI increases in sex chromosomal aberrations might be a result of non-random chromosomal positioning and defects in male gamete nuclear decondensation after the ooplasmic injections of non-acrosomally reacted spermatozoa [140]. In a recent study, Bonduelle et al. [141] carried out a medical follow-up study of 5-year-old ICSI children and compared the findings with a population of children born after natural conception. Growth assessed as sta-ture at follow-up was similar in the two groups despite a higher rate of preterm birth and low birthweight in the ICSI children. Common diseases and chronic illnesses occurred at similar rates in both groups. More ICSI children underwent surgical intervention and required other therapies.
3.8 Exogenous DNA and HIV transmission risks from use of ICSI procedures HIV infection or gamete contamination by exogenous DNA do not belong to genetic or epigenetic risks. However, they represent an issue of major concern in ICSI procedures. Transmission of viral elements, especially retroviruses which have the ability to integrate and transpose in the human genome, might represent a considerable risk. In more than 1 000 insemination cycles, artificial insemination involving HIV-seropositive males did not appear to be accompanied by transmitting the virus and 250 successful pregnancies were reported [142]. In addition, ICSI procedures using HIV-positive frozen semen samples have resulted in the generation of embryos free from the HIV virus [143_145]. Although in vitro preparation of semen samples by washing and gradient separation before the ICSI techniques are carried out appear to block the transmission of viruses, there is a potential risk of exogenous DNA transmission to the embryo. This hypothetical risk is based on studies in Rhesus Macaque monkeys showing that exogenous DNA bound to spermatozoa can be transferred by ICSI to the embryos and, thus, it might confer some new genetically transmitted characteristics [146]. Consequently, hypothetical binding of exogenous DNA on human spermatozoa processed for ICSI might alter the germline genetic constitution of the human offspring. A cautious manipulation of semen samples and use of strict safety procedures to exclude sources of DNA contamination during sperm manipulation are recommended in ICSI laboratories. For this reason in assisted reproduction programs, PGD procedures (using PCR) should be carried out in isolated facilities and thermal cyclers with UV decontaminators (that are separated from the ICSI laboratories) to eliminate the risk for transmission of exogenous DNA during ICSI procedures.
3.9 Genetic and epigenetic risks from the intraooplasmic injection of in vivo produced spermatids The introduction of the intracytoplasmic injection of spermatids or secondary spermatocytes as an alternative mode of therapy of non-obstructed azoospermic men who are negative for testicular foci of spermatozoa raised several concerns for probable genetic risks associated with the immaturity of the early haploid male gamete [16, 147_149]. The genetic risks of ooplasmic injections of human round spermatids might be a) inherent to the population of men this procedure is applied to (i.e. transferring chromosomal abnormalities/gene deletions to the offspring); or b) inherent to the procedure per se. The latter risks might be associated with abnormalities in the a) centrosomal components of the early haploid male gamete (defects in the reproducing element of the centrosome might cause zygotic spindle abnormalities after ooplasmic injections of spermatids) [16]; b) nuclear proteins; or c) spermatid oocyte-activating factor (i.e. the male gamete substance that triggers the cascade of ooplasmic events that result in the resumption of meiosis of the female gamete post-ooplasmic injections) [150_152]. In addition, it is particularly tempting to investigate in humans whether the process of genomic imprinting has been completed at the round spermatid stage [153]. This hypothesis has been evaluated in experimental mammals (Mus musculus) reproduced through ooplasmic injections of spermatids. The results have shown that there is no difference in the genomic imprinting establishment process between normally reproduced animals and animals generated from spermatids [154]. Studies in animals suggest that mouse genomic imprinting (Figure 2) is complete at/prior to the primary spermatocyte stage [155, 156]. The results of studies in our laboratory indicate that the genomic imprinting process in the rabbit and the rat has been completed at/before the round spermatid stage [157, 158]. It should be emphasized that even if genomic imprinting has not been completed at the round spermatid stage, the genomic imprinting process might be completed postfertilization (during early embryonic development) [16, 148, 159]. Regarding genomic imprinting abnormalities-related di-seases after ooplasmic injections of spermatids, there is no evidence today of imprinting defects in the offspring [16]. However, because methylation of some imprinted genes is supposed to occur during spermatogenesis or during early embryonic development [16, 159, 160], additional studies are necessary in order to evaluate the methylation status of genes in children delivered after ooplasmic injections of spermatids. Data on congenital and chromosomal abnormalities in children born after intracytoplasmic injection of spermatids are not sufficient to draw safe conclusions. Nevertheless, one report is alarming and indicates major abnormalities in children delivered after ooplasmic injections of spermatids [161]. Other studies on larger series did not detect an increased incidence of malformations after ooplasmic injections of spermatids [162_164]. However, considering that the number of human pregnancies achieved after ooplasmic injections of spermatids is limited, no definite conclusions can be drawn on the safety of ooplasmic injections of early haploid male gametes. Ejaculated round spermatids in the rat appear to have a lower reproductive capacity than testicular round spermatids [158]. This might be attributable to morphological defects in the ejaculated round spermatids (Figure 1). Another alteration the male gamete undergoes during spermiogenesis in vivo is the replacement of the nuclear histones (low disulphide bond proteins) by protamines (high disulphide bond proteins). Histones are protecting the early haploid male gamete DNA (within the cytoplasm of the oocyte) after ooplasmic injections. The presence of low disulphide bond proteins around the round spermatid DNA after round spermatid nuclei injections (ROSNI) or after round spermatid injections (ROSI) has been considered to be a factor responsible for the low outcome of these techniques. In contrast, post-ICSI, protamines are protecting the spermatozoal DNA within the ooplasm. In the case of ooplasmic injections of early spermatids, the survival of the injected spermatid DNA within the ooplasm might be detrimentally affected by the absence of protamines [16]. Post-ICSI, the resumption of meiosis of the female gamete depends on/is facilited by the presence of the oocyte-activating factor present in mouse, rabbit and human spermatozoa [150_152, 157, 165]. Defects in the expression/functionality of the oocyte activating factor after ooplasmic injections of early spermatids might account for their lower fertilization and pregnancy rates (comparatively with those after ICSI procedures). Although Kimura and Yanagimachi [150_152] and Sofikitis et al. [158] have shown that the oocyte-activating factor has not been expressed in mouse and rat round spermatids, respectively, several studies suggest that the oocyte activating factor has been expressed in the round spermatid in the human or the rabbit [16, 166_168]. Healthy offspring have been delivered after prede-condensed sperm or even spermatid head injections into the female pronuclei of preactivated rat oocytes [169]. The latter study might suggest that novel methods of assisted syngamy have been developed and such a technology in the future might have a role in cases of human ICSI failure as a result of lack of development of male pronucleus (post-ICSI) or inability of the male and female pronuclei to fuse.
3.10 Genetic risks after assisted reproduction techniques using in vitro generated male haploid germ cells Although induction of human meiosis and spermiogenesis in an in vitro culture system represents an attractive alternative solution for the therapeutic management of men who are positive for spermatogonia/spermatocytes but negative for haploid cells in their testes, the application of diploid germ cell in vitro culture technique might be limited by ethical considerations or safety-related factors. For instance, application of ooplasmic injections of human haploid cells generated in in vitro culture systems containing xenogeneic Sertoli cells [111, 164, 170] is susceptible to ethical considerations and risks regarding contamination of the human germ cells by animal viruses or animal molecules. Similarly, a major drawback for application of ooplasmic injections of haploid male gametes derived in in vitro co-culture systems of human diploid germ cells with supporting animal feeder somatic cells, such as Vero or STO cells, concerns the risks of transmitting infectious agents to the human germ cells [164]. The growth phase of Vero cells is usually achieved in the presence of newborn calf serum, which still poses the risk of virus or animal molecule transmission to the cultured human cells [171]. In addition, performance of assisted reproduction procedures using immature haploid germ cells derived or cultured in vitro is susceptible to genetic and epigenetic risks. Kimura et al. [156] attempted to induce both male meiotic divisions in vitro within the cytoplasm of oocytes injected with primary spermatocytes. They observed a high frequency of abnormalities in male meiotic chromosomal behavior when mouse primary spermatocytes were injected into the ooplasm of MII oocytes. It seems that most primary spermatocytes have not acquired the competence for normal chromosomal segregation within the ooplasm and/or that the ooplasm does not provide adequate factors required to segregate the spermatocyte chromosomes that are still synapsed. In humans, Sousa et al. [163] reported that most of the embryos, produced after ooplasmic injections of spermatids that had been generated in vitro, showed sex chromosomal abnormalities. The high abnormal genetic constitution of the derived human embryos might have been to the result of: a) a deficient male meiotic process in vitro; or b) the immature DNA-status of the in vitro generated haploid cells. Tesarik et al. [166_168] showed a very rapid progression of meiosis and/or spermiogenesis during in vitro culture of human primary spermatocytes and/or round spermatids, respectively. It is possible that the action of multiple checking mechanisms, which control/coordinate the male gamete morphogenetic and molecular transformations during spermatogenesis in vivo, cannot be completed (totally or partially) during the in vitro culture of spermatogenic cells. The overall result might be a high percentage of abnormal products of meiosis and/or spermiogenesis in in vitro culture systems. This is consistent with the fact that an increase in DNA degradation of round spermatids during in vitro culture has been observed [168]. Thus, it appears that the clinical employment of ooplasmic injections of in vitro derived haploid germ cells might be associated with genetic risks attributable to the completion of meiosis or a part of the spermiogenetic process under in vitro conditions.
3.11 Epigenetic risks related to assisted reproduction techniques Genomic imprinting abnormalities might also have an impact on assisted reproductive techniques in which spermatozoa are injected into oocytes. Only one copy (paternal or maternal) of an imprinted gene is active (Figure 2) and the other, the inactive one, is epigenetically "marked" by histone modification, cytosine methylation or both [172]. It has been shown that the mammalian primordial male germ cell genome undergoes extensive epigenetic reprogramming, namely demethylation (i.e. erasure of the previous imprint), to assure later at the gamete stage the establishment/consolidation of the maternal or the paternal imprint. Epigenetic marks originating from the parental cells must be erased at an early stage. Both copies of an imprinted gene are marked de novo during spermatogenesis according to the sex they originate from. After the consolidation of the new imprint, one of the two copies remains silent. After fertilization, imprinted genes maintain their methylation status and they escape the reprogramming (demethylation and reme-thylation) process. In contrast, it has been suggested that the methylation process in the unmethylated genes continues postfertilization [16, 159]. Alarming reports have recently raised concerns regarding the increased incidence of children with rare imprinting disorders, namely Angelman and Beckwith-Wiedemann syndromes (BWS), among children conceived by assisted reproduction. Two independent groups from USA and Europe have reported cases of Angelman syndrome conceived by ICSI techniques with sporadic imprinting defects [4, 173]. The mosaic methylation pattern detected in one of the patients and the absence of imprinting center mutations might support the evidence of a postzygotic epigenetic defect [174]. Furthermore, the analysis of chromosome 15 methylation pattern in a limited number of ICSI children (n = 92) did not show methylation abnormities [175]. BWS, a rare genetic condition (1/15 000), has also been reported to show a more frequent incidence among ICSI children [176_178]. It is worth mentioning that the study of DeBaun et al. [176] was prospective and identified an incidence of BWS equal to 4.6% among the children delivered after assisted reproduction techniques versus the background rate of 0.8% in the USA. Imprinting mutations of two BWS related genes were found in 5/6 children with BWS syndrome born after assisted reproduction [176]. The identification of Angelman syndrome and BWS syndrome among ICSI children indicate the need for additional prospective studies. In the above mentioned reports concerning BWS and AS patients, the epigenetic defect was found in the maternal allele suggesting that the abnormality might not be related to the spermatozoa used for ICSI. Whether or not imprinting defects are related to the culture conditions, media used to the hyperstimulation protocols or other epigenetic defects related to the development of male infertility but yet unidentified remains to be elucidated [179]. As we have recently mentioned [164], achievement of the induction of meiosis of male diploid germ cells and partial completion of spermiogenesis under in vitro conditions might not be accompanied by all the epigenetic modifications the male gamete normally undergoes during the respective stages of spermatogenesis under in vivo conditions. Additional epigenetic modifications, such as DNA methylation, genomic imprinting, RNA silencing and modification of histones, are important for the in vitro derived haploid male gamete nucleus in order to survive within the ooplasm and trigger the cascade of events that lead to normal embryonic development [174]. Acceleration of the cytoplasmic and nuclear maturation events that occur in vitro in cultured male germ cells might cause a disturbance of epigenetic reprogramming resulting in aberrant gene expression, abnormal phenotypic characteristics, and defects in the male gamete capacity to fertilize the oocyte and induce normal embryonic development. As we have emphasized in the above paragraphs, an important issue is whether genomic imprinting establishment has been completed in immature diploid or haploid male gametes. Kerjean et al. [180] showed that the methylation patterns of H19 and MEST/PEG1 genes are established as early as spermatogonial differentiation in humans. In contrast, Ariel et al. [181] showed that spermatogenesis-specific genes undergo late epigenetic reprogramming at the level of epididymis. Hajkova et al. [182] have shown that mouse PGC exhibit dynamic changes in epigenetic modifications between days 10.5 and 12.5 post coitum. PGC acquire genome-wide de novo methylation during early development and migration into the genital ridge. However, following their entry into the genital ridge there is a rapid erasure of DNA methylation of regions within imprinted and non-imprinted loci. Thus, there is an active demethylation process initiated upon the entry of PGC into the gonadal anlagen. The time of reprogramming of PGC is of paramount importance, because it ensures that germ cells in the males acquire a certain epigenetic state prior to the differentiation of the definitive male germ cells in which new parental imprints are then established [182]. Defects in the epigenetic reprogramming in any cultured (in vitro) immature diploid germ cell population might result in the inheritance of epimutations in the haploid cells generated from the culture of the immature germ cells. The fact that DNA methyltransferase is present in spermatids might be an argument against the hypothesis that genomic imprinting is complete at the round spermatid stage. Another hypothesis is that even if the genomic imprinting has not been completed at the round spermatid stage, the male gamete genomic imprinting might be completed after the transfer of immature haploid spermatogenic cells within the ooplasm [150, 151], or even during the early embryonic development [16, 159]. This hypothesis is supported by the fact that waves of DNA methylation have been shown during early embryonic development, the blastocyst stage and the time of implantation [159]. There are several studies providing evidence for the presence of activity of the DNA methyltransferase during early embryonic development [16, 159]. In addition, from a limited data available, it appears that the imprint establishment has been completed in humans by the time the spermatid stage is reached [154, 174]. Although most of the above studies tend to suggest that the genomic imprinting process in humans has been completed prior to the spermatid stage in vivo, it is unknown whether the rapidly proceeding meiosis and early spermiogenesis occurring under conditions present in in vitro culture systems allow the completion of genomic imprinting process within these relatively short periods. This is a question of clinical importance because abnormalities in the completion of genomic imprinting during in vitro gametogenesis may be manifested (postfertilization) as tumor susceptibility or/and tumorgenesis. There are epigenetic differences (Figure 2) between the parental genomes during the evolution of genomic imprinting in mammals. These epigenetic differences between the parental genomes are enhanced in the zygote by means of DNA demethylation of the paternal genome soon after fertilization, whereas the maternal genome shown de novo methylation [183]. Such opposite effects on the parental genomes within the same oocyte cytoplasm might be achieved by the differential binding of stored cytoplasmic factors to the parental genomes [184]. Arney et al. [184] have shown a preferential interaction of HP1beta protein with the maternal genome immediately after sperm entrance into the mouse oocyte. Paternal genome binding of HP1beta is only detected at the pronuclear stage. Considering that it is unknown whether oocytes at the two pronuclei plus se-cond polar body stage that have been fertilized by in vitro-generated human haploid male gametes (generated from the culture of human primary spermatocytes of men with primary testicular damage) [164, 185] show normal paternal genome-binding of HP1beta, it appears that the probability that ooplasmic injections of in vitro-derived early haploid male gametes being accompanied by epigenetic risks related to a lack of or abnormalities in the pattern of binding of HP1beta protein with the paternal genome cannot be ruled out. It should be emphasized that in the theoretical case of injecting an imprint-free immature male germ cell nucleus into an oocyte, fertilization might be anticipated but it should lead to embryonic lethality. Transplantation of imprint-free PGC nuclei into oocytes has resulted in embryonic lethality, partly as a result of abnormal extraembryonic tissues resulting from the inappropriate silence or activation of imprinted genes [186]. So far, imprinting during passage through at least some stages of spermatogenesis is essential because a male genome devoid of imprints cannot acquire all of them within a mature oocyte [186]. In addition to the above described epigenetic factors, defects in other epigenetic factors might contribute to the abnormal characteristics of embryos produced by ICSI procedures [16, 163]. Abnormalities/defects in the expression of oocyte-activating factor in spermatozoa (see above paragraphs) might result in defects in the capacity of the male gamete (after its entrance into the ooplasm) to activate the cascade of ooplasmic events that result in resumption of meiosis of the female gamete, fertilization and normal embryonic development. Furthermore, deficiency in the functionality of the reproducing element of the centrosome [187], or the presence of an abnormal number of centrioles in spermatozoa, might cause aberrant spindle formation after ICSI techniques resulting in abnormal embryonic development. Defects in the paternally inherited centrosomic components are known to represent a reason for ICSI failure (to induce appropriate embryonic development) after the entrance of the male gamete into the ooplasm [187]. In addition, Luetjens et al. [188] showed that abnormalities in the male gamete nucleus condensation could retard the sperm X chromosome decondensation resulting in embryonic aneuploidy through zygotic mitotic errors. Thus, we cannot rule out the probability that a) abnormalities in the nuclear condensation status of spermatozoa or b) abnormalities in the capacity of spermatozoa to decondense at an appropriate chronological order within the ooplasm (post-ICSI) might cause chromosomal abnormalities in the embryos.
3.12 Risks concerning transgenerational transmission of an acquired genetic or epigenetic defect Apart from the genetic and epigenetic risks already described (which are substantiated by the abnormalities found in the offspring of patients treated with assisted reproduction procedures), there are also other less obvious risks. These risks may be called "risks concerning transgenerational transmission of an acquired genetic or epigenetic defect" and are mainly of two types: a) those resulting from the action of aggressive cancer treatment on gametes with overall genetic and teratogenetic consequences; and b) those that are anticipated in the future generations of ICSI offspring and concern defects in tumor suppression genes and increased susceptibility of ICSI-children for tumor development. It has been reported that there is a higher incidence of retinoblastoma among children conceived after assisted reproduction technology [189]. Although male gamete DNA damage might be inevitable during cancer treatment (i.e. chemotherapy, radia-tion) there is no evidence today of increased frequency of genetic defects or congenital malformations among children (either naturally conceived or conceived after ICSI techniques) fathered by men who have undergone chemotherapy. Nevertheless, DNA breaks are induced by reactive oxygen species produced either by aggressive cancer therapy or during sperm preparation techniques for carrying out assisted reproductive technology or by microorganisms contaminating the lower genitourinary tract [190, 191]. Furthermore, DNA denaturation and fragmentation are strongly correlated with a decreased reproductive potential [192]. Fertilization of an oocyte (using ICSI techniques) with a DNA-damaged spermatozoon might be accompanied by a risk for a genetic disease in the offspring.
3.13 Risks related to mutations of genes regulating the spermiogenesis process The process of spermiogenesis is very sensitive to genetic alterations. Alterations in the expression of molecular agents in the testicular tissue as a result of defects in gene expression (null mutations, gene over-expression, exogenous gene expression and gene misexpression) could lead to a deficiency in the completion of specific steps of spermiogenesis. These defects in gene expression might result in spermatogenic arrest at the round spermatid stage or in the production of few spermatozoa with anatomical or functional defects. Although men with arrest at the round spermatid stage or oligozoospermic men with anatomically or functionally deficient spermatozoa do not have reproductive potential under in vivo conditions, ICSI procedures or ooplasmic injections of spermatids might offer the latter men the probability to father their own children. However, the bypassing via assisted reproductive technology of biological barriers related to defects in the spermiogenesis process is accompanied by risks for transferring gene defects to the male assisted reproductive technology offspring. The expression of phenotypic characteristics (i.e. defects in spermiogenesis) in the offspring (generated by assisted reproductive technology) depends on the chromosomal location of the respective mutated gene, the pattern of the inheritance of this gene and/or the pre-sence of any type of mutations/alterations in the expression of this gene in the mother's genotype. To emphasize the importance of mutations in genes regulating spermiogenesis, we are describing below some genes playing a role in the spermiogenesis process. Histone replacement by transition proteins (TP) and protamines during spermiogenesis might be affected by disruption of the Tarbp2 gene, resulting in infertility and oligospermia [193]. A partial or complete failure to synthesize the protamines results in delayed replacement of TP and the spermatids show abnormal nuclear morphogenesis, developmental arrest and degeneration [193]. Premature translation of Prm1 (pre-existing protamine 1) mRNA cause precocious condensation of spermatid nuclear DNA and abnormal head morphogenesis [194]. Successful interaction of mature protamine-2 with chromatin is required for displacement of TP2 [195]. Step-15 spermatids in Camk4-/- mice show a loss of protamine-2. These animals are characterized by prolonged retention of TP2. Mice lacking the major TP1 have been obtained after targeted deletion of the Tnp1 gene. Tnp1-/- mice show a normal sperm production quantitatively, but only 23% of the spermatozoa show any movement and most of these spermatozoa do not show forward progression [195,196]. In these animals, sperm heads with a blunted or bent tip are seen in 16% of epididymal spermatozoa, possibly generated by the abnormal chromatin condensation that could reduce the rigidity of the fine apex of the spermatozoon [195,196]. Tnp1 contains a cAMP-responsive element (CRE) that serves as a binding site for the CRE modulator (CREM). CREM is involved in the regulation of Tnp1 gene expression and human CREM protein is synthesized in steps 1_3 round spermatids. This might explain why a reduction in Crem expression and a lack of both CREM and TP1 have been shown in human spermatids arrested at step 3 [197]. Mice with deletion in Crem presented a spermatogenesis arrest at the round spermatid step [198]. Deficiencies in intratesticular molecular factors as a result of genetic defects affect the organization and reorganization of the cytoskeleton during spermiogenesis. Thus, homozygous c-ros knockout mice are sterile and the epididymal spermatozoa have bent tails and compromised flagellar vigour within the uterus [199]. Testicular haploid expression gene (THEG) is expressed in round and elongated spermatids. The molecular products of this gene appear to play a role in the spermiogenesis because abnormal or absent flagella in mice with THEG dysruption have been shown and might be to the result of an impairment of the assembly of cytoskeletal proteins such as the tubulins [200]. A specific block in spermiogenesis was observed in homozygous JunD-/- mice. A lack of molecular factors encoded by the latter gene results in an absence of flagella in spermatids in the lumen of the seminiferous tubules [201, 202]. The absence of JunD led to sperm flagellar growth impairment. Additional defects in sperm nuclear and cytoskeletal morphology, and in mitochondrial localization can be observed in nectin-null mutant mice. Nectin-2 is a component of cell-cell anchoring junctions, playing a role in the connection of the cytoskeletal elements of neighbouring cells. Thus, this molecular system participates in the regulation of cell shape and differentiation through signalling pathways [203]. Further interesting observations on the male gamete cytoskeleton are shown in the null mutant for the zinc-finger transcription factor Egr4. In the latter animals, the flagella is often fragmented, sharply kinked or have tightly coiled distal ends. Spermatozoa with heads that are either separated entirely or bent sharply back on the flagella are observed [202, 204]. In null mice for Sla12a2 gene (normally expressing the Na+-K+-2Cl- co-transporter), few spermatids are present but defects are striking when spermatids gradually acquire the features of spermatozoa [202]. Defects in the molecular system of Na+-K+-2Cl- co-transporter result in morphological abnormalities of spermatids. Spermatids show abnormalities in the cap-phase acrosomal vesicle and in the nuclear shape [205]. Other morphological abnormalities of the male gamete are the result of the lack of the factors that are normally expressed by the CsnK2a2 gene. CsnK2a2 could be a candidate globo-zoospermia gene. Mice with defects in the expression of the CsnK2a2 gene show abnormalities in spermatid nuclear morphogenesis. Further abnormalities are observed in the nuclear and acrosomal shape. Robertson et al. [206] have shown that deficiency in the production of aromatase enzyme cyp19 as a result of targeted disruption of the cyp19 gene in ArKO mice results in maturation arrest at early stages of spermiogenesis. Round spermatids do not complete elongation and spermiation. Furthermore, morphological defects in round spermatids are seen in tubules exhibiting spermiogenic arrest. Furthermore, abnormalities in the acrosomal structure are observed. Deficiency in the production of an epithelial, microtubule-associated protein due to defects in the expression of the E-MAP-115 gene results in abnormal shape and progressive degeneration in all condensed spermatids. Abnormalities in the microtubular manchette and in nuclear shape are also observed [202, 207]. Subnormal expression of the molecular products of the gene Tg737 that encodes the components of the raft protein complex, designated Polaris in the mouse and IFT88 in both Chlamydomonas and mouse, results in defective ciliogenesis and abnormalities in flagellar development in spermatids as well as asymmetry in left-right axis determination [208]. Polaris/IFT88 is detected in the manchette of mouse and rat spermatids. Intramanchette transport has the features of intraflagellar transport machinery. In addition, it facilitates nucleocytoplasmic exchange activities during spermiogenesis [208]. 3.14 Preimplantation genetic diagnosis (PGD)-biopsy techniques and risks Monogenic and chromosomal abnormalities can be diagnosed using genetic material obtained from polar bodies (PB), blastomeres or trophectoderm cells [209]. Couples who have had previous unsuccessful assisted reproductive trials or have a risk of transmitting to the offspring a genetic disorder related or unrelated to their infertility status might benefit from the application of PGD. PND is the most widely applied procedure, however, it is often followed by iatrogenic termination of a pregnancy associated with a fetal recessive or dominant disorder or with a fetus numerical or structural chromosomal abnormality [210, 211]. The clinical application of PGD has a number of limitations concerning: a) its diagnostic value; b) the availability of oocytes, zygotes or embryos for biopsy; and c) the implantation or pregnancy rates after the healthy embryo transfer [212]. Embryonic biopsy, as an invasive method, might also have risks related to the post-PGD embryonic development and, furthermore, to the health of newborns [22]. For instance, there is evidence that acid Tyrode solution (commonly used to carry out PB biopsy) affects the quality and the development of the embryos that have undergone biopsy, despite the fact that the aspiration of both PB does not cause a detrimental effect on the cleavage of the zygote [213]. Currently, the usage of acid Tyrode solution is gradually being replaced by laser drilling of the zona pellucida and, thus, the utilization of chemical substances is substituted by the use of a high-energy beam. Studies comparing the two methods have been in favor of the laser drilling in terms of implantation and pregnancy rates post-biopsy [214]. Analysis of data regarding the carrying out of PGD has indicated that the PB biopsy is not used as often as the blastomere biopsy and is practically limited to cases of oocyte selection (to carry out ICSI) in female carriers of chromosomal translocations [211, 212, 215]. To carry out blastomere biopsy, at least one blastomere is aspirated from all day-3 embryos. A second (additional to the first) blastomere biopsy offers reassurance of the validity and reliability of the diagnosis, although it increases the workload of the clinical PGD procedure. The implantation and pregnancy rates related to the two cell (blastomeres) biopsy are similar to the one cell biopsy. Thus, it appears that the aspiration of a second blastomere does not have a detrimental affect on further embryonic development [216]. In addition, there is evidence proving that transfer of blastocysts (on day 5) that have been generated from embryos that had undergone biopsy on day-3 embryos does not compromise the implantation process [217]. Regarding the potential risks arising from the blastomere biopsy at the 6_8 cell stage embryo, there is concern originating from the evidence that the X chromosome inactivation process is initiated at this developmental stage [218]. Biopsy of one or two blastomeres from a limited pool of cells might disrupt the 50%/50% ratio of the random X chromosome inactivation balance [219]. Pediatric evaluation of children born after ICSI plus PGD did not show significant differences compared with children born after the use of ICSI trials [22, 220]. Biopsy of the trophectoderm is an alternative method to the blastomere biopsy with a limited experience to date [209]. However, it should be emphasized that blastocysts provide a sufficient amount of genetic material for reliable diagnosis. In addition, blastocysts that have undergone biopsy have an acceptable capacity for implantation [221]. Couples with infertility related genetic abnormalities might benefit from the use of PGD. For these couples the balance between risks and benefits supports the role of PGD to select genetically competent embryos and to avoid the PND during the pregnancy period.
4 Guidelines and conclusions
According to the guidelines suggested by a group of clinical and research experts from 12 national scientific societies, there are two types of genetic tests for ICSI candidates: a) the recommended tests; and b) the optional tests according to the clinical indications [222]. The genetic profiles of both members of each couple participating in assisted reproductive technology programs has to be carefully assessed and proper genetic counseling and a basic genetic evaluation (Table 1) will assist all couples to make informed decisions. The highly recommended diagnostic tests for infertile males participating in assisted reproductive programs include the karyotype, microdeletions of the Y chromosome and the CFTR mutation analysis. Additional genetic tests for KAL 1 mutations, androgen receptor, 5 a-reductase 2, hemoglobinopathies and sperm-aneuploidy analysis might be additionally suggested for selected subpopulations of infertile males (Tables 1, 2). Data regarding thousands of children evaluated in independent studies, pooled data form surveys of world results and the ESHRE ICSI task force, as well, show that the proportions of the most common congenital abnormalities in children delivered after ICSI techniques are not significantly different compared with those in the general population, with the exception of hypospadias [223_226]. Reservations concern the definitions of major and minor abnormalities, and the abnormalities that should be evaluated in a long run, such as deficiencies in mental development. The genetic profiles and constitution of gametes from males treated with ICSI are variable, however, it appears that a relationship exists between the severity of the spermatogenic impairment and the chromosomal defects in the spermatozoa (either testicular or ejaculated samples) [48, 114]. In addition, germline genetic defects in spermatogenesis have to be taken into consideration when ICSI is suggested [65]. Genetic counseling by experienced scientists should emphasize that even mild or isolated phenotypic defects in the father may lead to more severe and clinically important abnormalities in the offspring. Non-obstructed azoospermic men with complete deletions of AZFa or AZFb region of the Y chromosome (Table 2, Figure 3) should not be advised to undergo testicular biopsy. The development of ICSI as a widely applied and prominent reproductive technology has intensified the need for thorough evaluation and laboratory investigations towards two directions. The first direction is the follow-up of children derived by ICSI techniques and the second target is the analysis/study of the genetic causes underlying male infertility. Results from both directions might give rise to conclusions regarding the pathogenesis and the role of male infertility/primary testicular damage in the generation of male gamete (and subsequently embryonic) chromosomal abnormalities. Today, ICSI might be additionally applied as a result of other indications, for example PGD. PGD can assist the genetic safety of ICSI. The effective collaboration of fertility specialists and geneticists, taken together with the introduction of thorough genetic evaluation in assisted reproduction programs, is essential to reduce the genetic risks from the application of modern reproductive technology.
References
1 Qian Y, Feng T, Chen J, Cai LB, Liu JY, Mao YD, et al. Fertilization of in vitro matured human oocytes by intracytoplasmic sperm injection (ICSI) using ejaculated and testicular spermatozoa. Asian J Androl 2005; 7: 39_43. 2 Johnson MD. Genetic risks of intracytoplasmic sperm injection in the treatment of male infertility: recommendations for genetic counselling and screening. Fertil Steril 1998; 70: 397_411. 3 Thompson JG, Kind KL, Roberts CT, Robertson SA, Robinson JS. Epigenetic risks related to assisted reproductive technologies: short- and long-term consequences for the health of children conceived through assisted reproduction technology: more reason for caution? Hum Reprod 2002; 17: 2783_6. 4 Cox GF, Burger J, Lip V, Mau UA, Sperling K, Wu BL, et al. Intracytoplasmic sperm injection may increase the risk of imprinting defects. Am J Hum Genet 2002; 71: 162_4. 5 Anway MD, Li Y, Ravindranath N, Dym M, Griswold MD. Expression of testicular germ cell genes identified by differential display analysis. J Androl 2003; 24: 173_84. 6 Anway MD, Ravindranath N, Dym M, Griswold MD. Identification of a murine testis complementary DNA encoding a homolog to human A-kinase anchoring protein-associated sperm protein. Biol Reprod 2002; 66: 1755_61. 7 Anway MD, Cupp AS, Uzumcu M, Skinner MK. Epigenetic transgenerational actions of endocrine disruptors and male fertility. Science 2005; 308: 1466_9. 8 Anway MD, Skinner MK. Epigenetic transgenerational actions of endocrine disruptors. Endocrinology 2006; 147(Suppl.): S43_S49. 9 Matzuk MM, Lamb DJ. Genetic dissection of mammalian fertility pathways. Nat Cell Biol 2002; 4 (Suppl): 41_9. 10 Tung YJ, Luetjens CM, Wistuba J, Xu EY, Reijo Pera RA, Gromoll J. Evolutionary comparison of the reproductive genes, DAZL and BOULE, in primates with and without DAZ. Dev Genes Evol 2006; 216: 158_68. 11 Agulnik AI, Zharkikh A, Boettger-Tong H, Bourgeron T, McElreavey K, Bishop EC. Evolution of the DAZ gene family suggests tha Y-linked DAZ plays little, or a limited, role in spermatogenesis but underlines a recent African origin for human populations. Hum Mol Gen 1998; 7: 1371_7. 12 Reynolds N, Cooke HJ. Role of the DAZ genes in male fertility. Reprod Biomed Online 2005; 10: 72_80. 13 Meschede D, Horst J. The molecular genetics of male infertility. Mol Hum Reprod 1997; 3: 419_30. 14 Diemer T, Desjardins C. Developmental and genetic disorders in spermatogenesis. Hum Reprod Update 1999; 5: 120_40. 15 Kupker W, Schwinger E, Hiort O, Ludwig M, Nikolettos N, Schlegel PN, et al. Genetics of male subfertility: consequences for the clinical work-up. Hum Reprod 1999; 14 (Suppl. 1): 24_37. 16 Sofikitis N, Miyagawa I, Yamamoto Y, Loutradis D, Mantza-vinos T, Tarlatzis V. Micro- and macro-consequences of ooplasmic injections of early haploid male gametes. Hum Reprod Update 1998; 4: 197_212 17 Patrizio P, Leonard DGB. Mutations of the cystic fibrosis gene and congenital absence of the vas deferens. In: McElreavey K, editor, The Genetic Basis of Male Infertility. Berlin: Springer Verlag, 2000: 175_86. 18 De Braekeleer M, Ferec C. Mutations in the cystic fibrosis gene in men with congenital bilateral absence of vas deferens. Mol Hum Reprod 1996; 2: 669-77. 19 Cuppens H, Lin W, Jaspers M, Costes B, Teng H, Vankeer-berghen A, et al. Polyvariant mutant cystic fibrosis transmembrane conductance regulator genes. The polymorphic (Tg)m locus explains the partial penetrance of the T5 polymorphism as a disease mutation. J Clin Invest 1998; 101: 487_96. 20 Kiesewetter S, Macek M Jr, Davis C, Curristin SM, Chu CS, Graham C, et al. A mutation in CFTR produces different phenotypes depending on chromosomal background. Nat Genet 1993; 5: 274_8. 21 Liebaers I, Sermon K, Staessen C, Joris H, Lissens W, Van Assche E, et al. Clinical experience with preimplantation genetic diagnosis and intracytoplasmic sperm injection. Hum Reprod 1998; 13 (Suppl. 1): 186_95. 22 Vandervors M, Staessen C, Sermon K, De Vos A, Van de Velde H, Van Assche E, et al. The Brussels' experience of more than 5 years of clinical preimplantation genetic diagnosis. Hum Reprod Update 2000; 6: 364_73. 23 Josserand RN, Bey-Omar F, Rollet J, Lejeune H, Boggio D, Durand DV, et al. Cystic fibrosis phenotype evaluation and paternity outcome in 50 males with congenital bilateral absence of vas deferens. Hum Reprod 2001; 16: 2093_7. 24 Nieschlag E. Classification of andrological disorders. In Nieschlag E, Behre HM, editors, Andrology: Male Reproductive Health and Dysfunction. Berlin: Springer-Verlag, 1997: 81_3. 25 Nieschlag E, Behre HM, Meschede D. Disorders at the testicular level. In Nieschlag E, Behre HM, editors, Andrology: Male Reproductive Health and Dysfunction. Berlin: Springer-Verlag, 1997: 133_59. 26 Bartoloni L, Blouin JL, Pan Y, Gehrig C, Maiti AK, Scamuffa N, et al. Mutations in the DNAH11 (axonemal heavy chain dynein type 11) gene cause one form of situs inversus totalis and most likely primary ciliary dyskinesia. Proc Natl Acad Sci USA 2002; 99: 10282_6. 27 Zamboni L. The ultrastructural pathology of the spermatozoon as a cause of infertility: the role of electron microscopy in the evaluation of semen quality. Fertil Steril 1987; 48: 711_34. 28 Neugebauer DC, Neuwinger J, Jockenhovel F, Nieschlag E. `9 + 0' axoneme in spermatozoa and some nasal cilia of a patient with totally immotile spermatozoa associated with thickened sheath and short midpiece. Hum Reprod 1990; 5: 981_6. 29 Schill WB. Some disturbances of acrosomal development and function in human spermatozoa. Hum Reprod 1991; 6: 969_78. 30 Pirrello O, Machev N, Schimdt F, Terriou P, Menezo Y, Xiville S. Search for mutations involved in human globozoospermia. Hum Reprod 2005; 20: 1314_8. 31 Behre HM, Nieschlag E, Behre HM, editors, Andrology: Male Reproductive Health and Dysfunction. Berlin: Springer-Verlag, 1997: 115_29. 32 Kalantaridou SN, Chrousos GP. Monogenic disorders of puberty. J Clin Endocrinol Metab 2002; 87: 2481_94. 33 Willems PJ. Dynamic mutations hit double figures. Nat Genet 1994; 8: 213_5. 34 McLean HE, Warne GL, Zajac JD. Defects of androgen receptor function: from sex reversal to motor neuron disease. Mol Cell Endocrinol 1995; 112: 133_41. 35 Mifsud A, Sim CK, Boettger-Tong H, Moreira S, Lamb DJ, Lipshultz LI, et al. Trinucleotide (CAG) repeat polymorphisms in the androgen receptor gene: molecular markers of risk for male infertility. Fertil Steril 2001; 75: 275_ 81. 36 Georgiou I, Sermon K, Lissens W, De Vos A, Platteau P, Lolis D, Van Steirteghem A, Liebaers I, et al. Preimplantation genetic diagnosis for spinal and bulbar muscular atrophy (SBMA). Hum Genet 2001; 108: 494_8. 37 Hortas ML, Castilla JA, Gil MT, Molina J, Garrido ML, Morell M, et al. Decreased sperm function of patients with myotonic muscular dystrophy. Hum Reprod 2000; 15: 445_8. 38 Pan H, Li YY, Li TC, Tsai WT, Li SY, Hsiao KM. Increased (CTG/CAG)(n) lengths in myotonic dystrophy type 1 and Machado-Joseph disease genes in idiopathic azoospermia patients. Hum Reprod 2002; 17: 1578_83. 39 Dean NL, Phillips SJ, Chan P, Tan SL, Ao A. Reported relationship between increased CTG repeat lengths in myotonic dystrophy and azoospermia. Hum Reprod 2002; 17: 3003_4. 40 Sermon K, De Vos A, Van de Velde H, Seneca S, Lissens W, Joris H, et al. Fluorescent PCR and automated fragment analysis for the clinical application of preimplantation genetic diagnosis of myotonic dystrophy (Steinert's disease). Mol Hum Reprod 1998; 4: 791_6. 41 Sherman S. Epidemiology. In Hagerman RJ, Hagerman PJ, editors, Fragile X syndrome. Baltimore and London: The Johns Hopkins University Press, 2002: 136_68. 42 Nistal M, Martinez-Garcia F, Regadera J, Cobo P, Paniagua R. Macro-orchidism: Light and electron microscopic study of four cases. Hum Pathol 1992; 23: 1011_8. 43 Tamanini F, Willemsen R, van Unen L, Bontekoe C, Galjaard H, Oostra BA, et al. Differential expression of FMR1, FXR1 and FXR2 proteins in human brain and testis. Hum Mol Genet 1997; 6: 1315_22. 44 Platteau P, Sermon K, Seneca S, VanSteirtheim A, Devroey P, Liebaers I. Preimplantation genetic diagnosis for fragile Xa syndrome: difficult but not impossible. Hum Reprod 2002; 17: 2807_12. 45 Allingham-Hawkins DJ, Babul-Hirji R, Chitayat D, Holden JJ, Yang KT, Lee C, et al. Fragile X premutation is a significant risk factor for premature ovarian failure: the International Collaborative POF in Fragile X study—preliminary data. Am J Med Genet 1999; 83: 651_2. 46 Sermon K, Seneca S, Vanderfaeillie A, Lissens W, Joris H, Vandervorst M, et al. Preimplantation diagnosis for fragile X syndrome based on the detection of the non-expanded paternal and maternal CGG. Prenat Diagn 1999; 19: 1223_30. 47 Van Assche E, Bonduelle M, Tournaye H, Joris H, Verheyen G, Devroey P, et al. Cytogenetics of infertile men. In: Steirteghem AV, Devroey P, Liebaers I, editors, Genetics and Assisted Human Conception. Hum Reprod 1996; 11 (Suppl. 4): 1_24. 48 Palermo GD, Colombero LT, Hariprashad JJ, Schlegel PN, Rosenwaks Z. Chromosomal analysis of epididymal and testicular sperm in azoospermic patients undergoing ICSI. Hum Reprod 2002; 17: 570_5. 49 Mateizel I, Verheyen G, Van Assche E, Tournaye H, Liebaers I, Van Steirteghem A. FISH analysis of chromosome X,Y and 18 abnormalities in testicular sperm from azoospermic patients. Hum Reprod 2002; 17: 2249_57. 50 Devroey P, Van Steirteghem A. A review of ten years experience of ICSI. Human Reprod Update 2004; 10: 19_28. 51 Bonduelle M, Van Assche E, Joris H, Keymolen K, Devroey P, Van Steirteghem A, et al. Prenatal testing in ICSI pregnancies: incidence of chromosomal anomalies in 1586 karyotypes and relation to sperm parameters. Hum Reprod 2002; 17: 2600_14. 52 Yamamoto Y, Sofikitis N, Mio Y, Loutradis D, Kaponis A, Miyagawa I. Morphometric and cytogenetic characteristics of testicular germ cells and Sertoli cell secretory function in men with non-mosaic Kleinefelters syndrome. Hum Reprod 2002; 17: 886_96. 53 Chandley AC, Edmond P, Christie S, Gowans L, Fletcher J, Frackiewicz A, et al. Cytogenetics and infertility in man. I. Karyotype and seminal analysis: results of a five-year survey of men attending a subfertility clinic. Ann Hum Genet 1975; 39: 231_54. 54 Elliott DJ, Cooke HJ. The molecular genetics of male infertility. Bioessays 1997; 19: 801_9. 55 Olesen C, Hansen C, Bendsen E, Byskov AG, Schwinger E, Lopez-Pajares I, et al. Identification of human candidate genes for male infertility by digital differential display. Mol Hum Reprod 2001; 7: 11_20. 56 Bache I, Assche EV, Cingoz S, Bugge M, Tumer Z, Hjorth M, et al. An excess of chromosome 1 breakpoints in male infertility. Eur J Hum Genet 2004; 12: 993_1000. 57 Guichaoua MR, Speed RM, Luciani JM, Delafontaine D, Chandley AC. Infertility in human males with autosomal translocations. II. 95 Meiotic studies in three reciprocal rearrangements, one showing tertiary monosomy in a 45-chromosome individual and his father. Cytogenet Cell Genet 1992; 60: 96_101. 58 Siffroi JP, Benzacken B, Straub B, Le Bourhis C, North MO, Curotti G, et al. Assisted reproductive technology and complex chromosomal rearrangements: the limits of ICSI. Mol Hum Reprod 1997;3: 847_51. 59 Staessen C, Van Steirteghem AC. The chromosomal constitution of embryos developing from abnormally fertilized oocytes after intracytoplasmic sperm injection and conventional in-vitro fertilization. Hum Reprod 1997; 12: 321_7. 60 Meschede D, Lemcke B, Exeler JR, De Geyter C, Behre HM, Nieschlag E, et al. Chromosome abnormalities in 447 couples undergoing intracytoplasmic sperm injection—prevalence, types, sex distribution and reproductive relevance. Hum Reprod 1998; 13: 576_82. 61 Scriven PN, Flinter FA, Braude PR, Ogilvie CM. Robertsonian translocations—reproductive risks and indications for preimplantation genetic diagnosis. Hum Reprod 2001; 16: 2267_73. 62 Munne S, Fung J, Cassel MJ, Marquez C, Weier HU. Preimplantation genetic analysis of translocations: case-specific probes for interphase cell analysis. Hum Genet 1998; 102: 663_74. 63 Van Assche E, Staessen C, Vegetti W, Bonduelle M, Vandervorst M, Van Steirteghem A, et al. Preimplantation genetic diagnosis and sperm analysis by fluorescence in-situ hybridization for the most common reciprocal translocation t(11;22). Mol Hum Reprod 1999; 5: 682_90. 64 Scriven PN, O'Mahony F, Bickerstaff H, Yeong CT, Braude P, Mackie Ogilvie C. Clinical pregnancy following blastomere biopsy and PGD for a reciprocal translocation carrier: analysis of meiotic outcomes and embryo quality in two IVF cycles. Prenat Diagn 2000; 20: 587_92. 65 Yamamoto Y, Sofikitis N, Kaponis A, Georgiou J, Giannakis D, Mamoulakis CH, et al. Use of a highly sensitive quantitative telomerase assay in intracytoplasmic sperm injection programmes for the treatment of 47,XXY non-mosaic Klinefelter men. Andrologia 2002; 34: 218_26. 66 Foresta C, Galeazzi C, Bettella A, Stella M, Scandellari C. High incidence of sex chromosome aneuploidies in two patients with Klinefelter syndrome. J Clin Endocrinol Metab 1998; 83: 203_5. 67 Tournaye H, Liu J, Nagy Z, Joris H, Wisanto A, Bonduelle M, et al. Intracytoplasmic sperm injection (ICSI): the Brussels experience. Reprod Fertil Dev 1995; 7: 278_9. 68 Staessen C, Coonen E, Van Assche E, Tournaye H, Joris H, Devroey P, et al. Preimplantation diagnosis for X and Y normality in embryos from three Klinefelter patients. Hum Reprod 1996; 11: 1650_3. 69 Palermo GD, Schlegel PN, Sills ES, Veeck LL, Zaninovic N, Menendez S, et al. Births after intracytoplasmic injection of sperm obtained by testicular extraction from men with nonmosaic Klinefelter's syndrome. N Engl J Med 1998; 338: 588_90. 70 Ron-El R, Strassburger D, Gelman-Kohan S, Friedler S, Raziel A, Appelman Z. A 47,XXY fetus conceived after ICSI of spermatozoa from a patient with non-mosaic Klinefelter's syndrome: case report. Hum Reprod 2000; 8: 1804_6. 71 Akashi T, Fuse H, Kojima Y, Hayashi M, Honda S. Birth after intracytoplasmic sperm injection of ejaculated spermatozoa from a man with mosaic Klinefelter's syndrome. Asian J Androl 2005; 7: 217_20. 72 Bettella A, Ferlin A, Menegazzo M, Ferigo M, Tavolini IM, Bassi PF, et al. Testicular fine needle aspiration as a diagnostic tool in non-obstructive azoospermia. Asian J Androl 2005; 7: 289_94. 73 Chevret E, Rousseaux S, Monteil M, Usson Y, Cozzi J, Pelletier R, et al. Increased incidence of hyperhaploid 24,XY spermatozoa detected by three-colour FISH in a 46,XY/47,XXY male. Hum Genet 1996; 97: 171_5. 74 Chandley AC. Infertility. In: Rimoin DL, Connor JM, Pyeritz RE, editors, Principles and Practice of Medical Genetics. 3rd ed. New York: Churchill Livingstone Inc., 1997:667_75. 75 Blanco J, Rubio C, Simon C, Egozcue J, Vidal F. Increased incidence of disomic sperm nuclei in a 47,XYY male assessed by fluorescent in situ hybridization. Hum Genet 1997; 99: 413_6. 76 Madan K. Balanced structural changes involving the human X: effect on sexual phenotype. Hum Genet 1983;63:216_21. 77 Kalz-Fuller B, Sleegers E, Schwanitz G, Schubert R. Charac-terisation, phenotypic manifestations and X-inactivation pattern in 14 patients with X-autosome translocations. Clin Genet 1999; 55: 352_66. 78 Solari AJ, Rahn IM, Ferreyra ME, Carballo MA. The beha-vior of sex chromosomes in two human X-autosome trans-locations: failure of extensive X-inactivation spreading. Biocell 2001; 25: 155_66. 79 Handel MA, Hunt PA. Sex chromosome pairing and activity during mammalian meiosis. Bioessays 1992; 14: 817_22. 80 Jamieson RV, Tam PP, Gardiner-Garden M. X-chromosome activity: impact of imprinting and chromatin structure. Int J Dev Biol 1996; 40: 1065_80. 81 Solari AJ. The behavior of the XY pair in mammals. Int Rev Cytol 1974; 38: 273_317. 82 Meschede D, Froster UG, Bergmann M, Nieschlag E. Fami-lial pericentric inversion of chromosome 1(p34q23) and male infertility with stage specific spermatogenic arrest. J Med Genet 1994; 31: 573_5. 83 Navarro J, Benet J, Martorell MR, Templado C, Egozcue J. Segregation analysis in a man heterozygous for a pericentric inversion of chromosome 7(p13;q36) by sperm chromosome studies. Am J Hum Genet 1993; 53, 214_9. 84 Gabriel-Robez O, Ratomponirina C, Croquette M, Couturier J, Rumpler Y. Synaptonemal complexes in a subfertile man with a pericentric inversion in chromosome 21. Heterosynapsis without previous homosynapsis. Cytogenet Cell Genet 1988; 48: 84_7. 85 ISLAT (Institute for Science Law, and Technology) Working Group. ART into science: regulation of fertility techniques. Science 1998; 281: 651_2. 86 Katagiri Y, Neri QV, Takeuchi T, Schlegel PN, Megid WA, Kent-First M, et al. Y chromosome assessment and its implications for the development of ICSI children. Reprod Biomed Online 2004; 8: 307_18. 87 Zeng S, Patil RS, Yankowitz J. Prenatal detection of mosaic trisomy 1q due to an unbalanced translocation in one fetus of a twin pregnancy following in vitro fertilization: a postzygotic error. Am J Med Genet 2003; 120: 464_9. 88 Delobel B, Djlelati R, Gabriel-Robez O, Croquette MF, Rousseaux-Prevost R, Rousseaux J, et al. Y-autoxome translocation and infertility: usefulness of moleculsr, cytogenetic and meiotic studies. Hum Genet 1998; 102: 98_102. 89 Pinho MJ, Neves R, Costa P, Ferras C, Sousa M, Alves C, et al. Unique t(Y;1)(q12;q12) reciprocal translocation with loss of the heterochromatic region of chromosome 1 in a male with azoospermia due to meiotic arrest: a case report. Hum Reprod 2005; 20: 689_96. 90 Chandley AC. Chromosomes. In Hargreave TJ, editor, Male Infertility, 2nd ed., New York: Springer-Verlag, 1994; 149_64. 91 Yoshida A, Nakahori Y, Kuroki Y, Motoyama M, Araki Y, Miura K, et al. Dicentric Y chromosome in an azoospermic male. Mol Hum Reprod 1997; 3: 709_12. 92 Sasagawa I, Ishigooka M, Kato T, Hayami S, Hashimoto T, Nakada T. Dicentric Y chromosome without evidence of mosaicism in an azoospermic male. Scand J Urol Nephrol 1996; 30: 75_6. 93 Van Landuyt L, Lissens W, Stouffs K, Tournaye H, Liebaers I, Van Steirteghem A. Validation of a simple Yq deletion screening program in an ICSI candidate population. Mol Hum Reprod 2000; 6: 291_7. 94 Huynh T, Mollard R, Trounson A. Selected genetic factors associated with male infertility. Hum Reprod 2002; 8: 183_98. Simoni M, Bakker E, Krausz C. EAA/EMQN best practice guidelines for molecular diagnosis of y-chromosomal microdeletions. State of the art 2004. Int J Androl 2004; 27: 240_9. 96 Vogt PH, Edelmann A, Kirsch S, Henegariu O, Hirschmann P, Kiesewetter F, et al. Human Y chromosome azoospermia factors (AZF) mapped to different subregions in Yq11. Hum Mol Genet 1996; 5: 933_43. 97 Krausz C, Forti G, McElreavey K. The Y chromosome and male fertility and infertility. Int J Androl 2003; 26: 70-5. 98 Luetjens CM, Gromoll J, Engelhardt M, Von Eckardstein S, Bergmann M, Nieschlag E, et al. Manifestation of the Y-chromosomal deletion in human testis: a morphometrical and immunoistochemical evaluation. Hum Reprod 2002; 17: 2258_66. 99 Giachini C, Guarducci E, Longepied G, Degl'Innocenti S, Becherini L, Forti G, et al. The gr/gr deletion. A new genetic test in male infertility? J Med Genet 2005; 42: 497_502. 100 Teng YN, Lin YM, Lin YH, Tsao SY, Hsu CC, Lin SJ, et al. Association of a single-nucleotide polymorphism of the deleted-in-azoospermia-like gene with susceptibility to spermatogenic failure. J Clin Endocrinol Metab 2002; 87: 5258_64. 101 Schlegel PN. The Y chromosome. Reprod Biomed Online 2002; 5: 22_5. 102 Pryor JL, Kent-First M, Muallem A, Van Bergen AH, Nolten WE, Meisner L, et al. Microdeletions in the Y chromosome of infertile men. N Engl J Med 1997; 336: 534_9. 103 Kuhnert B, Gromoll J, Kostova E, Tschanter P, Luetjens CM, Simoni M, et al. Case report: natural transmission of an AZFc Y-chromosome microdeletion from father to his sons. Hum Reprod 2004; 19: 886_8. 104 Rolf C, Gromoll J, Simoni M, Nieschlag E. Natural transmission of a partial AZFb deletion of the Y chromosome over three generations: case report. Hum Reprod 2002; 17: 2267_71. 105 Kamishcke A, Gromoll J, Simoni M, Behre HM, Nieschlag E. Transmission of a Y chromosomal deletion involving the deleted in azoospermia (DAZ) and chromodomain (CDY1) genes from father to son through intracytoplasmic sperm injection. Hum Reprod 1999; 14: 2320_2 106 Kent-First MG, Kol S, Muallem A, Ofir R, Manor D, Blazer S, et al. The incidence and possible relevance of Y-linked microdeletions in babies born after intracytoplasmic sperm injection and their infertile fathers. Mol Hum Reprod 1996; 2: 943_50. 107 Krausz C, Quintana-Murci L, McElreavey K. Prognostic value of Y deletion analysis: what is the clinical prognostic value of Y chromosome microdeletion analysis? Hum Reprod 2000; 15: 1431_4. 108 Vogt PH. Human chromosome deletions in Yq11, AZF candidate genes and male infertility: history and update. Mol Hum Reprod 1998; 4: 739_44. 109 Maurer B, Simoni M. Y chromosome microdeletion screening in infertile men. J Endocrinol Invest 2000; 23: 664_70. 110 Patsalis PC, Sismani C, Quintana-Murci L, Taleb-Bekkouche F, Krausz C, McElreavey K. Effects of transmission of Y chromosome AZFc deletions. Lancet 2002; 360: 1222_4. 111 Sofikitis N, Kaponis A, Mio Y, Makredimas D, Giannakis D, Yamamoto Y, et al. Germ cell transplantation: a review and progress report on ICSI from spermatozoa generated in xenogeneic testes. Hum Reprod Update 2003; 9: 291_307. 112 Silber SJ, Repping S. Transmission of male infertility to future generations: lessons from the Y chromosome. Hum Reprod Update 2002; 8: 217_29. 113 Faddy MJ, Silber SJ, Gosden RG. Intra-cytoplasmic sperm injection and infertility. Nat Genet 2001; 29: 131. 114 Pang MG, Hoegerman SF, Cuticchia AJ, Moon SY, Doncel GF, Acosta AA, et al. Detection of aneyploidy for chromosomes 4, 6, 7, 8, 9, 10, 11, 12, 13, 17, 18, 21, X and Y by fluorescence in-situ hybridization in spermatozoa from nine patients with oligoasthenospermia undergoing intracytoplasmic sperm injection. Hum Reprod 1999; 14: 1266_73. 115 Vegetti W, Van Assche E, Frias A, Verheyen G, Bianchi MM, Bonduelle M, et al. Correlation between semen parameters and sperm aneuploidy rates investigated by fluorescence in situ hybridization in infertile men. Hum Reprod 2000; 15: 351_65. 116 Huang WJ, Lamb DJ, Kim ED, de Lara J, Lin WW, Lipshultz LI, et al. Germ cell nondisjuction in testes biopsies of men with idiopathic infertility. Am J Hum Genet 1999; 64: 1638_45. 117 Viville S, Warter S, Meyer JM, Wittemer C, Loriot M, Mollard R, et al. Histological and genetic analysis and risk assessment for chromosomal aberration after ICSI for patients presenting with CBAVD. Hum Reprod 2000; 15: 1613_8. 118 Platteau P, Staessen C, Michiels A, Tournaye H, Van Steirteghem A, Liebaers I, et al. Comparison of the aneuploidy frequency in embryos derived from testicular sperm extraction in obstructive and non-obstructive azoospermic men. Hum Reprod 2004; 19: 1570_4. 119 Piasecka M, Kawiak J. Sperm mitochondria of patients with normal sperm motility and with asthenozoospermia: morphological and functional study. Folia Histochem Cytobiol 2003; 41: 125_39. 120 Alosilla Fonttis A, Napolitano R, Tomas MA. Successful ICSI in a case of severe asthenozoospermia due to 93% non-specific axonemal alterations and 90% abnormal or absent mitochondrial sheaths. Reprod Biomed Online 2002; 5: 270_2. 121 Danan C, Sternberg D, Van Steirteghem A, Cazeneuve C, Duquesnoy P, Besmond C, et al. Evaluation of parental mitochondrial inheritance in neonates born after intracytoplasmic sperm injection. Am J Hum Genet 1999; 65: 463_73. 122 Bonduelle M, Legein J, Derde MP, Buysse A, Schietecatte J, Wisanto A, et al. Comparative follow-up study of 130 children born after ICSI and 130 children born after IVF. Hum Reprod 1995; 10: 3327_31. 123 Bonduelle M, Liebaers I, Deketelaere V, Derde MP, Camus M, Devroey P, et al. Neonatal data on a cohort of 2 889 infants born after ICSI (1991-1999) and of 2 995 infants born after IVF (1983-1999). Hum Reprod 2002; 17: 671_94. 124 In't Veld P, Brandenburg H, Verhoeff A, Dhont M, Los F. Sex chromosomal abnormalities and intracytoplasmic sperm injection. Lancet 1995; 346: 773. 125 Van Steirteghem A, Bonduelle M, Liebaers I, Devroey P. Children born after assisted reproductive technology. Am J Perinatol 2002; 19: 59_65. 126 Bonduelle M, Camus M, De Vos A, Staessen C, Tournaye H, Van Assche E, et al. Seven years of intracytoplasmic sperm injection and follow-up of 1987 subsequent children. Hum Reprod 1999; 14 (Suppl 1): 243_64. 127 Wennerholm UB, Bergh C, Hamberger L, Lundin K, Nilsson L, Wikland M, et al. Incidence of congenital malformations in children born after ICSI. Hum Reprod 2000;15: 944_8. 128 Ericson A, Kallen B. Congenital malformations in infants born after IVF: a population-based study. Hum Reprod 2001; 16: 504_9. 129 Peschka B, Leygraaf J, Van der Ven K, Montag M, Schartmann B, Schubert R, et al. Type and frequency of chromosome aberrations in 781 couples undergoing intracytoplasmic sperm injection. Hum Reprod 1999; 14: 2257_63. 130 Bonduelle M, Joris H, Hofmans K, Liebaers I, Van Steirteghem A. Mental development of 201 ICSI children at 2 years of age. Lancet 1998; 351: 1553. 131 Bonduelle M, Aytoz A, Van Assche E, Devroey P, Liebaers I, Van Steirteghem A. Incidence of chromosomal aberrations in children born after assisted reproduction through intracytoplasmic sperm injection. Hum Reprod 1998; 13: 781_2. 132 Bonduelle M, Wilikens A, Buysse A. A follow-up study of children born after intracytoplasmic sperm injection (ICSI) with epididymal and testicular spermatozoa and after replacement of cryopreserved embryos obtained after ICSI. Hum Reprod 1998; 13 (suppl): 196_207. 133 Bonduelle M, Wennerholm UB, Loft A, Tarlatzis BC, Peters C, Henriet S, et al. A multi-centre cohort study of the physical health of 5-year-old children conceived after intracytoplasmic sperm injection, in vitro fertilization and natural conception. Hum Reprod 2005; 20: 413_9. 134 Ponjaert-Kristoffersen I, Bonduelle M, Barnes J, Nekkebroeck J, Loft A, Wennerholm UB, et al. International collaborative study of intracytoplasmic sperm injection-conceived, in vitro fertilization-conceived, and naturally conceived 5-year-old child outcomes: cognitive and motor assessments. Pediatrics 2005; 115: 283_9. 135 Antoni K, Hamori M. Distribution of fetal malformations and chromosomal disorders in 1920 ICSI newborns between 1993 and 2000. Hum Reprod 2001; 16: 39. 136 Egozcue S, Blanco J, Vendrell JM, Garcia F, Veiga A, Aran B, et al. Human male infertility: chromosome anomalies, meiotic disorders, abnormal spermatozoa and recurrent abortion. Hum Reprod Update 2000; 6: 93_105. 137 Vernaeve V, Tournaye H, Osmanagaoglu K, Verheyen G, Van Steirteghem A, Devroey P. Intracytoplasmic sperm injection with testicular spermatozoa is less successful in men with nonobstructive azoospermia than in men with obstructive azoospermia. Fertil Steril 2003; 79: 529_33. 138 Aytoz A, Camus M, Tournaye H, Bonduelle M, Van Steirteghem A, Devroey P. Outcome of pregnancies after intracytoplasmic sperm injection and the effect of sperm origin and quality on this outcome. Fertil Steril 1998; 70: 500_5. 139 Shi Q, Martin RH. Aneuploidy in human spermatozoa: FISH analysis in men with constitutional chromosomal abnormalities, and in infertile men. Reproduction 2001; 121: 655_66. 140 Hewitson L, Dominko T, Takahashi D, Martinovich C, Ramalho-Santos J, Sutovsky P, et al. Unique checkpoints during the first cycle of fertilization after intracytoplasmic sperm injection in rhesus monkeys. Nat Med 1999; 5: 431_3. 141 Bonduelle M, Bergh C, Niklasson A, Palermo GD, Wennerholm UB; Collaborative Study Group of Brussels, Gothenburg and New York. Medical follow-up study of 5-year-old ICSI children. Reprod Biomed Online 2004; 9: 91_101. 142 Semprini A. Fiore S, Pardi G. Reproductive counseling for HIV discordant couples. Lancet 1997; 349: 1401_2. 143 Marina S, Marina F, Alcolea R, Nadal J, Exposito R, Huguet J. Pregnancy following intracytoplasmic sperm injection from an HIV-1 seropositive man. Hum Reprod 1998;13: 3247_9. 144 Kiessling AA. Expression of human immunodeficiency virus long terminal repeat-coupled genes in early cleaving embryos. J Reprod Immunol 1998; 41: 95_104. 145 Loutradis D, Drakakis P, Kallianidis K, Patsoula E, Bletsa R, Michalas S. Birth of two infants who were seronegative for human immunodeficiency virus type 1 (HIV-1) after intracytoplasmic injection of sperm from HIV-1-seropositive men. Fertil Steril 2001; 75: 210_2. 146 Chan AW, Luetjens CM, Dominko T, Ramalho-Santos J, Simerly CR, Hewitson L, et al. Foreign DNA transmission by ICSI: injection of spermatozoa bound with exogenous DNA results in embryonic GFP expression and live Rhesus monkeys births. Mol Hum Reprod 2000; 6: 26_33. 147 Tesarik J, Mendoza C, Testart J. Viable embryos from injections of round spermatids into oocytes. N Engl J Med 1995; 333: 525. 148 Sofikitis NV, Yamamoto Y, Miyagawa I, Mekras G, Mio Y, Toda T, et al. Ooplasmic injection of elongating spermatids for the treatment of non-obstructive azoospermia. Hum Reprod 1998;13:709_14. 149 Sofikitis N, Mantzavinos T, Loutradis D, Yamamoto Y, Tarlatzis V, Miyagawa I. Ooplasmic injections of secondary spermatocytes for non-obstructive azoospermia. Lancet 1998; 18: 351: 1177_8. 150 Kimura Y, Yanagimachi R. Mouse oocytes injected with testicular spermatozoa or round spermatids can develop into normal offspring. Development 1995; 121: 2397_405. 151 Kimura Y, Yanagimachi R. Development of normal mice from oocytes injected with secondary spermatocyte nuclei. Biol Reprod 1995; 53: 855_62. 152 Kimura Y, Yanagimachi R. Intracytoplasmic sperm injection in the mouse. Biol Reprod 1995; 52: 709_20. 153 Stewart CL, Pedersen R, Rotwein P, Bestor T, Rastan S, Hastie N, et al.; NIEHS/EPA Workshops. Genomic imprinting. Reprod Toxicol 1997; 11: 309_16. 154 Shamanski FL, Kimura Y, Lavoir MC, Pedersen RA, Yanagimachi R. Status of genomic imprinting in mouse spermatids. Hum Reprod 1999; 14: 1050_6. 155 Ogura A, Matsuda J, Suzuki O. Zygote-constructing ability of spermatogenic cells in mammals. Tanpakushitsu Kakusan Koso 1998; 43: 522_9. 156 Kimura Y, Tateno H, Handel MA, Yanagimachi R. Factors affecting meiotic and developmental competence of primary spermatocyte nuclei injected into mouse oocytes. Biol Reprod 1998; 59: 871_7. 157 Sofikitis NV, Miyagawa I, Agapitos E, Pasyianos P, Toda T, Hellstrom WJ, et al. Reproductive capacity of the nucleus of the male gamete after completion of meiosis. J Assist Reprod Genet 1994; 11: 335_41. 158 Sofikitis N, Ono K, Yamamoto Y, Papadopoulos H, Miyagawa I. Influence of the male reproductive tract on the reproductive potential of round spermatids abnormally released from the seminiferous epithelium. Hum Reprod 1999; 14: 1998_2006. 159 Fishel S, Aslam I, Tesarik J. Spermatid conception: a stage too early, or a time too soon? Hum Reprod 1996; 11: 1371_5. 160 Bestor TH. Cytosine methylation and the unequal developmental potentials of the oocyte and sperm genomes. Am J Hum Genet 1998; 62: 1269_73. 161 Zech H, Vanderzwalmen P, Prapas Y, Lejeune B, Duba E, Schoysman R. Congenital malformations after intracytoplasmic injection of spermatids. Hum Reprod 2000; 15: 969_71. 162 Sousa M, Fernandes S, Barros A. Prognostic factors for successful testicle spermatid recover. Mol Cell Endocrinol 2000; 166: 37_43. 163 Sousa M, Cremades N, Alves C, Silva J, Barros A. Developmental potential of human spermatogenic cells co-cultured with Sertoli cells. Hum Reprod 2002; 17: 161_72. 164 Sofikitis N, Pappas E, Kawatani A, Baltogiannis D, Loutradis D, Kanakas N, et al. Efforts to create an artificial testis: culture systems of male germ cells under biochemical conditions resembling the seminiferous tubular biochemical environment. Hum Reprod Update 2005; 11: 229_59. 165 Tesarik J, Sousa M, Testart J. Human oocyte activation after intracytoplasmic sperm injection. Hum Reprod 1994; 9: 511_8. 166 Tesarik J, Guido M, Mendoza C, Greco E. Human spermatogenesis in vitro: respective effects of follicle stimulating hormone and testosterone on meiosis, spermiogenesis, and Sertoli cell apoptosis. J Clin Endocrinol Metab 1998; 83: 4467_73. 167 Tesarik J, Sousa M, Greco E, Mendoza C. Spermatids as gametes: indications and limitations. Hum Reprod1998; 3 (Suppl. 13): 89_107. 168 Tesarik J, Greco E, Cohen-Bacrie P, Mendoza C. Germ cell apoptosis in men with complete and incomplete spermiogenesis failure. Mol Hum Reprod 1998; 4: 757_62. 169 Sofikitis N, Baltogiannis D, Takenaka M, Tsoukanelis K, Tsambalas S, Yamamoto Y, et al. Pre-decondensed sperm head injections into female pronuclei result in chromosomal mingling, zygotic cleavage, and adequate embryonic and fetal development up to delivery of healthy offspring: a novel method of assisted syngamy. Andrologia 2004; 36: 291_304. 170 Kawamura H, Kaponis A, Tasos A, Giannakis D, Miyagawa I, Sofikitis N. Rat Sertoli cells can support human meiosis. Hum Reprod 2003; 18(Suppl 1): 78 Presented in 19th Annual Meeting of ESHRE, Madrid, Spain. 171 Cremades N, Sousa M, Bernabeu R, Barros A. Developmental potential of elongating and elongated spermatids obtained after in-vitro maturation of isolated round spermatids. Hum Reprod 2001; 16: 1938_44. 172 Reik W, Dean W, Walter J. Epigenetic reprogramming in mammalian development. Science 2001; 293: 1089_93. 173 Orstavik KH, Eiklid K, van der Hagen CB, Spetalen S, Kierulf K, Skjeldal O, et al. Another case of imprinting defect in a girl with angelman syndrome who was conceived by intracytoplasmic semen injection. Am J Hum Genet 2003; 72: 218_9. 174 De Rycke M, Liebaers I, Van Steirteghem A. Epigenetic risks related to assisted reproductive technologies: risk analysis and epigenetic inheritance. Hum Reprod 2002; 17: 2487_94. 175 Manning M, Lissens W, Bonduelle M, Camus M, De Rijcke M, Liebaers I, et al. Study of DNA-methylation patterns at chromosome 15q11-q13 in children born after ICSI reveals no imprinting defects. Mol Hum Reprod 2000; 6: 1049_53. 176 DeBaun MR, Niemitz EL, Feinberg AP. Association of in vitro fertilization with Beckwith-Wiedemann syndrome and epigenetic alterations of LIT1 and H19. Am J Hum Genet 2003; 72: 156_60. 177 Maher ER, Brueton LA, Bowdin SC, Luharia A, Cooper W, Cole TR, et al. Beckwith-Wiedemann syndrome and assisted reproduction technology (ART). J Med Genet 2003; 40: 62_4. 178 Gicquel C, Gaston V, Mandelbaum J, Siffroi JP, Flahault A, Le Bouc Y. In vitro fertilization may increase the risk of Beckwith-Wiedemann syndrome related to the abnormal imprinting of the KCN1OT gene. Am J Hum Genet 2003; 72: 338_41. 179 Niemitz EL, Feinberg AP. Epigenetics and assisted reproductive technology: a call for investigation. Am J Hum Genet 2004; 74: 599_609. 180 Kerjean A, Dupont JM, Vasseur C, Le Tessier D, Cuisset L, Paldi A, et al. Establishment of the paternal methylation imprint of the human H19 and MEST/PEG1 genes during spermatogenesis. Hum Mol Genet 2000; 9: 2183_7. 181 Ariel M, Cedar H, McCarrey J. Developmental changes in methylation of spermatogenesis-specific genes include reprogramming in the epididymis. Nat Genet 1994; 7: 59_63. 182 Hajkova P, Erhardt S, Lane N, Haaf T, El-Maarri O, Reik W, et al. Epigenetic reprogramming in mouse primordial germ cells. Mech Dev 2002; 117: 15_23. 183 Mayer W, Niveleau A, Walter J, Fundele R, Haaf T. Deme-thylation of the zygotic paternal genome. Nature 2000; 403: 501_2. 184 Arney KL, Bao S, Bannister AJ, Kouzarides T, Surani MA. Histone methylation defines epigenetic asymmetry in the mouse zygote. Int J Dev Biol 2002; 46: 317_20. 185 Tesarik J, Bahceci M, Ozcan C, Greco E, Mendoza C. Restoration of fertility by in-vitro spermatogenesis. Lancet 1999; 353: 555_6. 186 Ferguson-Smith AC, Surani MA. Imprinting and the epigenetic asymmetry between parental genomes. Science 2001; 293: 1086_9. 187 Schatten G. The centrosome and its mode of inheritance: the reduction of centrosome during gametogenesis and its restoration during fertilization. Dev Biol 1994; 165: 299_335. 188 Luetjens C, Payne C, Schatten G. Non-random chromosome positioning in human sperm and sex chromosomal anomalies following intracytoplasmic sperm injection. Lancet 1999; 353: 1240. 189 Moll AC, Imhof SM, Cruysberg JR, Schouten-van Meeteren AY, Boers M, van Leeuwen FE. Incidence of retinoblastoma in children born after in-vitro feritilization. Lancet 2003; 361: 273_4. 190 Twigg J, Irvine DS, Houston P, Fulton N, Michael L, Aitken RJ. Iatrogenic DNA damage induced in human spermatozoa during sperm preparation: protective significance of seminal plasma. Mol Hum Reprod 1998; 4: 439_45. 191 Potts JM, Sharma R, Pasqualotto F, Nelson D, Hall G, Agarwal A. Association of ureaplasma urealyticum with abnormal reactive oxygen species levels and absence of leukocytospermia. J Urol 2000; 163: 1775_8. 192 Zini A, Bielecki R, Phang D, Zenzes MT. Correlations between two markers of sperm DNA integrity, DNA denatura-tion and DNA fragmentation, in fertile and infertile men. Fertil Steril 2001; 75: 674_7. 193 Zhong J, Peters AH, Lee K, Braun RE. A double-stranded RNA binding protein required for activation of repressed messages in mammalian germ cells. Nat Genet 1999; 22: 171_4. 194 Lee K, Haugen HS, Clegg CH, Braun RE. Premature translation of protamine-1 mRNA causes precocious nuclear condensation and arrests spermatid differentiation in mice. Proc Natl Acad Sci 1995; 92: 12451_5. 195 Yu YE, Zhang Y, Unni E, Shirley CR, Deng JM, Russell LD, et al. Abnormal spermatogenesis and reduced fertility in transition nuclear protein 1-deficient mice. Proc Natl Acad Sci USA 2000; 97: 4683_8. 196 Zhao M, Shirley CR, Yu YE, Mohapatra B, Zhang Y, Unni E, et al. Targeted disruption of the transition protein 2 gene affects sperm chromatin structure and reduces fertility in mice. Mol Cell Biol. 2001; 21: 7243_55. 197 Steger K, Klonisch T, Gavenis K, Behr R, Schaller V, Drabent B, et al. Round spermatids show normal testis-specific H1t but reduced cAMP-responsive element modulator and transition protein 1 expression in men with round-spermatid maturation arrest. J Androl 1999; 20: 747_54. 198 De Cesare D, Fimia GM, Sassone-Corsi P. Signaling routes to CREM and CREB: plasticity in transcriptional activation. Trends Biochem Sci 1999; 24: 281_5. 199 Yeung CH, Wagenfeld A, Nieschlag E, Cooper TG. The cause of infertility of male c-ros tyrosine kinase receptor knockout mice. Biol Reprod 2000; 63: 612_8 200 Yanaka N, Kobayashi K, Wakimoto K, Yamada E, Imahie H, Imai Y, et al. Insertional mutation of the murine kisimo locus caused a defect in spermatogenesis. J Biol Chem 2000; 275: 14791_4. 201 Thepot D, Weitzman JB, Barra J, Segretain D, Stinnakre MG, Babinet C, et al. Targeted disruption of the murine junD gene results in multiple defects in male reproductive function.. Development 2000; 127: 143_53. 202 Escalier D. Impact of genetic engineering on the understanding of spermatogenesis. Hum Reprod Update 2001; 7: 191_210. 203 Bouchard MJ, Dong Y, McDermott BM Jr, Lam DH, Brown KR, Shelanski M, et al. Defects in nuclear and cytoskeletal morphology and mitochondrial localization in spermatozoa of mice lacking nectin-2, a component of cell-cell adherens junctions. Mol Cell Biol 2000; 20: 2865_73. 204 Tourtellotte WG, Nagarajan R, Auyeung A, Mueller C, Mibrandt J. Infertility associated with incomplete spermatogenic arrest and oligozoospermia in Egr4-deficient mice. Development 1999; 126: 5061_71. 205 Pace AJ, Lee E, Athirakul K, Coffman TM, O'Brien DA, Koller BH. Failure of spermatogenesis in mouse lines deficient in the Na(+)-K(+)-2Cl(-) cotransporter. J Clin Invest 2000; 105: 441_50. 206 Robertson KM, O'Donnell L, Jones ME, Meachem SJ, Boon WC, Fisher CR, et al. Impairment of spermatogenesis in mice lacking a functional aromatase (cyp 19) gene. Proc Natl Acad Sci 1999; 6: 7986_91. 207 Komada M, McLean DJ, Griswold MD, Russell MD, Soriano P. E-MAP-115, encoding a microtubule-associated protein, is a retinoic acid-inducible gene required for spermatogenesis. Genes Dev 2000; 14: 1332_42. 208 Kierszenbaum AL. Intramanchette transport (IMT): managing the making of the spermatid head, centrosome, and tail. Mol Reprod Dev 2002; 63: 1_4. 209 Sermon K. Current concepts in preimplantation genetic diagnosis (PGD): a molecular biologist's view. Hum Reprod Update 2002; 8: 11_20. 210 Kanavakis E, Traeger-Synodinos J. Preimplantation genetic diagnosis in clinical practice. J Med Genet 2002; 39: 6_11. 211 ESHRE PGD Consortium Steering Committee. ESHRE Preimplantation Genetic Diagnosis Consortium: data collection III (May 2001). Hum Reprod 2002; 17: 233_46. 212 Sermon K, Moutou C, Harper J, Geraedts J, Scriven P, Wilton L, et al. ESHRE PGD Consortium data collection IV: May_December 2001. Hum Reprod 2005; 20: 19_34 213 Cieslak J, Ivakhnenko V, Wolf G, Sheleg S, Verlinsky Y. Three-dimensional partial zona dissection for preimplantation genetic diagnosis and assisted hatching. Fertil Steril 1999; 71: 308_13. 214 ESHRE PGD Consortium Steering Committee ESHRE Preimplantation Genetic Diagnosis (PGD) Consortium: data collection II. Hum Reprod 2000; 15: 2673_83. 215 Durban M, Benet J, Boada M, Fernandez E, Calafell JM, Lailla JM, et al. PGD in female carriers of balanced Robertsonian and reciprocal translocations by first polar body analysis. Hum Reprod Update 2001; 7: 591_602. 216 Van de Velde H, De Vos A, Sermon K, Staessen C, De Rycke M, Van Assche E, Lissens W, et al. Embryo implantation after biopsy of one or two cells from cleavage-stage embryos with a view to preimplantation genetic diagnosis. Prenat Diagn 2000; 20: 1030_7. 217 Palmer GA, Traeger-Synodinos J, Davies S, Tzetis M, Vrettou C, Mastrominas M, et al. Pregnancies following blastocyst stage transfer in PGD cycles at risk for beta-thalassaemic hemoglobinopathies. Hum Reprod 2002; 17: 25_31. 218 Lanasa MC, Hogge WA, Kubik C, Blancato J, Hoffman EP. Highly skewed X-chromosome inactivation is associated with idiopathic recurrent spontaneous abortion. Am J Hum Genet 1999; 65: 252_4. 219 Okamoto I, Otte AP, Allis CD, Reinberg D, Heard E. Epigenetic dynamics of imprinted X inactivation during early mouse development. Science 2004; 303: 644_9. 220 Strom CM, Levin R, Strom S, Masciangelo C, Kuliev A, Verlinsky Y. Neonatal outcome of preimplantation genetic diagnosis by polar body removal: the first 109 infants. Pediatrics 2000; 106: 650_3. 221 Sermon K, Van Steirteghem A, Liebaers I. Preimplantation genetic diagnosis. Lancet 2004; 363: 1633_41. 222 Foresta C, Ferlin A, Gianaroli L, Dallapiccola B. Guidelines for the appropriate use of genetic tests in infertile couples. Eur J Hum Genet 2002; 10: 303_12. 223 Chandley AC, Hargreave TB. Genetic anomaly and ICSI. Hum Reprod 1996; 11: 930_2. 224 Martin RH. The risk of chromosomal abnormalities following ICSI. Hum Reprod 1996; 11: 924_5. 225 Persson JW, Peters GB, Saunders DM. Is ICSI associated with risks of genetic disease? Implications for counselling practice and research. Hum Reprod 1996; 11: 921_4. 226 Davis TL, Yang GJ, McCarrey JR, Bartolomei MS. The H19 methylation imprint is erased and re-established differentially on the parental alleles during male germ cell development. Hum Mol Genet 22; 9: 2885_94. |
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