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Male Reproductive Tract |
Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, Texas 77030
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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male reproductive tract, sperm, sperm maturation, sperm motility and transport, testis
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Our understanding of the autosomal genetic regulation of male gonadal development has been accelerated through genetic engineering in mice (for review, see [2–4]). Single-, double-, and triple-gene knock-out, knock-in, gene-trapping, and transgenic studies have all been used in reverse-genetic approaches to identify the role of known autosomal and sex-linked genes in male infertility (for review, see [4–6]). In addition, the application of forward genetics in mice (a phenotype-driven approach) is currently being used to extend the biological information gained from traditional transgenic approaches and to identify new alleles at a given locus. For example, combining high-efficiency mutagenesis using N-ethyl-N-nitrosourea (ENU) and deletion analysis, the spermatogenic alleles associated with the quaking locus quakingviable (qkv) on mouse chromosome 17 and the juvenile development and fertility (jdf2) locus on mouse chromosome 7 were identified [7–9].
With respect to specific chromosomes and/or chromosomal regions involved in genetic regulation of male fertility, the Y chromosome, the t-complex on the proximal end of mouse chromosome 17, and autosomal translocations associated with 6p and 6q have been implicated in humans and in animal models [10–17]. In particular, recent studies concerning the long arm of human chromosome Y (Yq) have revealed approximately 15 gene families [18], many of which have been shown to play critical roles in male germ cell and gonadal development. Mouse chromosome 11 has previously been shown to contain many important genetic loci for male and female fertility. These include the spontaneous mouse mutant germ cell deficient (gcd), which manifests as infertility in both males and females and maps to 11A2–3 [19, 20]; the knock-out of the ubiquitin-conjugating enzyme 2b (Ube2b), which results in a block in gametogenesis [5]; the knock-out of growth differentiation factor-9 (gdf-9), which results in an early block in ovarian folliculogenesis [21]; and the knock-out allele of the testicular isoform of angiotensin-converting enzyme (Ace-T) which results in complete male infertility because of abnormalities in germ cell angiotensin-converting enzyme expression [22].
Recent technical advances are now enabling us to use whole-autosome genetic engineering to create precisely defined deletion and balancer chromosomes to ascribe functional information to genomic regions in vivo [8, 23, 24]. In the first study, to our knowledge, of its kind in mice, we used a precisely engineered 24-cM balancer chromosome on mouse chromosome 11 that encompasses the region between Trp53 (39 cM) and Wnt3 (63 cM) to identify new genomic loci associated with hematopoiesis, craniofacial and cardiac development, and fertility [25]. In the present study, we describe the characteristics of a three-member, noncomplementing male infertility mutant class identified from this screen.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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C57BL/6J males, obtained from the Jackson Laboratory, Bar Harbor, ME, were injected with three weekly 100-mg/kg doses of ENU according to a protocol outlined previously [26]. After recovering fertility, the males were bred into the inversion-balancer scheme also outlined previously [25]. In brief, mutagenized C57BL/6J male mice were crossed with female mice carrying a single copy of Inv(11)8BrdTrp53-Wnt3 [23]. The informative G1 "carrier" mice (because they "carry" one copy of the balancer chromosome on mouse chromosome 11) were crossed with Inv(11)8Brd/Re. Only the mice carrying the balancer (yellow ears and tail) and the mutagenized chromosome 11 (noncurly-haired coat) were informative. The informative G2 carrier mice from a single G1 female were intercrossed to produce G3 offspring. The viable G3 offspring were then classified as the recessive mutant class, which have two copies of the ENU-mutagenized chromosome 11 and, therefore, do not have yellow ears and tail (these mice are called homozygous mutant "testers"), or as the carrier class that carry one copy of the balancer chromosome (which gives the mice yellow ears and tail) and one copy of the ENU-mutagenized chromosome (these mice are called heterozygous mutant "carriers"). The G3 tester class mice were analyzed using the fertility screen described below. All investigations were conducted in accordance with the Guide for the Care and Use of Laboratory Animals (Copyright 1996, National Academy of Sciences).
Fertility Screen
Male and female tester mice from a single pedigree at 8–10 wk of age were housed in mating cages for 12 wk. If no progeny were born within the 12-wk period, the mating pair was separated and the male tester was out-crossed with a known-to-be-fertile female for an additional 12 wk. Conversely, the female tester was out-crossed with a male known to be fertile for an additional 12 wk. From the second mating, it could be determined whether the source of infertility for the pedigree was male factor, female factor, or both. To confirm inheritance and the location of the mutation within Trp53 and Wnt3 on mouse chromosome 11, the carrier G3 siblings were either inter-crossed to recover tester- and carrier-class mice or back-crossed to Inv(11)8Brd/Re and the resulting carrier class progeny inter-crossed to recover tester- and carrier-class mice.
Epididymal Sperm Counts
Sperm counts, testis histology, testis weight, and transmission-electron microscopy (TEM) were performed at 7 wk of age. Male mice were anesthetized with Isoflo (isoflurane USP; Abbott Laboratories, North Chicago, IL) before cervical dislocation. External genitalia were examined for any morphological abnormalities before making a vertical incision in the abdominal cavity. Testicular descent and size of the seminal vesicles were noted. The testis and epididymis were revealed by gently pushing on the testicular sacs. The caudal epididymis was carefully separated from the remainder of the urogenital tract and placed in 2 ml of PBS prewarmed to 37°C. Following 10 min at 37°C, the cauda was roughly chopped and removed from the dish before viewing the sperm using an inverted light microscope (WILD Heerbrugg, Heerbrugg, Switzerland) at 400x magnification. Sperm counts were performed using a hemocytometer (Hausser Scientific, Horsham, PA).
Testis Histology, In Situ Cell Death, and TEM
Testis tissue for histology and detection of in situ cell death was removed and immersed briefly in Bouin fixative (VWR, West Chester, PA) before weighing and further fixation in Bouin solution for 24 h at room temperature on a platform shaker. Following fixation, testes were embedded in paraffin, sectioned (thickness, 5 µm), and stained with hematoxylin.
Apoptotic cell death was examined in both male carrier and tester mice from the Infertile 4 (Inf4) Infertile 8 (Inf8), and Infertile 9 (Inf9) pedigrees (so named because of the order in which they were recovered from the fertility screen) at 7 wk of age using the In Situ Cell Death Detection Kit (Roche Molecular Biochemicals, Mannheim, Germany) according to manufacturer's instructions. The single addition to the protocol involved incubating the sections for 30 min at room temperature with 20 µg/ml of proteinase K in 10 mM Tris/HCl (pH 7.4) under a glass coverslip before addition of the terminal deoxynucleotidyl transferase (TdT)-mediated dUTP nick end labeling (TUNEL) reaction mixture. All reactions were conducted under glass coverslips.
The TEM was performed on 1 mm3 of testis extracted from carrier and tester male mice of Inf4, Inf8, and Inf9 pedigrees. Tissue was fixed with 2.5% glutaraldehyde in 0.1 M phosphate buffer (pH 6.9) for 2 h at room temperature before postfixation in 2% osmium tetroxide in the same buffer for 1 h. Samples were dehydrated in ethanol and embedded in Maraglas 655 (Electron Microscopy Services, Fort Washington, PA) after propylene oxide treatment. Thick sections (1 µm) were stained with toluidine blue; thin sections (60 nm) were stained with uranyl acetate and lead citrate.
Sperm Motility
The caudal epididymis was removed and placed in a 200-µl droplet of Hepes-buffered sperm motility media (111 mM NaCl/2 mM KCl/1.2 mM MgSO4·7H2O/0.36 mM Na2HPO4/5.6 mM glucose/1.1 mM sodium pyruvate/50 mM Hepes/10 mM NaHCO3/0.1 mM phenol red) [27] containing 0.5% Fraction V fatty acid-free BSA (Sigma-Aldrich, St Louis, MO) at 37°C under mineral oil (Sigma-Aldrich). Osmolality of the sperm motility media was measured to be between 300 and 310 mOsm. The cauda was torn using 22-guage needles and removed from the droplet. Sperm from carrier males were diluted 1:20, whereas sperm from mutant tester males were assessed undiluted. A total of 15 µl of sperm plus media from each animal were loaded onto a Cell-Vu Counting Chamber (Millennium Sciences Corp., New York, NY) prewarmed to 37°C. Motility was assessed using an inverted light microscope (WILD Heerbrugg) by counting the number of motile sperm with forward progression, motile sperm without forward progression, and nonmotile sperm in a given field-of-view, with a total of 200 sperm counted per animal and a minimum of five animals analyzed per group.
Sperm Morphology
Air-dried smears were prepared from sperm extracted in PBS. The smears were stained with hematoxylin and examined using a Zeiss inverted light microscope (Carl Zeiss, Inc., Oberkochen, Germany) at 400x magnification. Head and tail morphology were determined independently for each mouse with at least 100–200 sperm assessed for each sample. Head morphologies were categorized as normal, club shaped, or tail-less, whereas tail morphologies were classified as normal, abnormal midpiece, abnormal principal piece, short, and headless [28].
Complementation Testing
We mated tester females from Inf9 with a carrier male from Inf4 and Inf8, and we mated a carrier male from Inf4 with a carrier female from Inf8. As a result, the progeny from these matings were heterozygous for the balancer and the infertile mutation inherited from the mother (carriers) and heterozygous for the infertile mutation inherited from the mother and the infertile mutation inherited from the father (testers). The testers and carriers were analyzed at 7 wk by sperm count, testis histology, testis weight, and sperm motility.
Statistics
Significance (P < 0.05) was calculated by performing an F-test to determine variance, followed by a two-tailed t-test comparing testers and carriers in a single pedigree. All results are expressed as the mean ± SD.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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A total of 184 pedigrees were screened for fertility at G3, resulting in the identification of six putative male and two putative female infertile pedigrees [25]. In the present analysis, we examined the phenotypic class categorized as infertile with reduced epididymal sperm count and qualitatively nonmotile epididymal sperm. As mentioned, we have named these three independently isolated mutant pedigrees Inf4, Inf8, and Inf9 because of the order in which they were recovered from the screen.
Sperm Count Was Significantly Reduced in MaleTester Mice
The average epididymal sperm count was significantly reduced in tester mice from Inf4 (17.8 ± 14.2 x 106 in carriers vs. 0.2 ± 0.2 x 106 in testers; P < 0.01), Inf8 (5.6 ± 2.2 x 106 in carriers vs. 0.3 ± 0.6 x 106 in testers; P < 0.001), and Inf9 (5.9 ± 3.0 x 106 in carriers vs. 0.9 ± 1.2 x 106 in testers; P < 0.001) pedigrees. Only in Inf4 did this result in a significant decrease of testis weight (114.7 ± 13.3 mg in carriers vs. 83.7 ± 16.3 mg in testers) (Table 1). Dissection of males in all pedigrees revealed phenotypically normal external genitalia, seminal vesicle size, and testicular descent (results not shown).
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Testis Histology and In Situ Cell Death
Given that the epididymal sperm count was significantly reduced, the stage of spermatogenic arrest was examined by histology (Fig. 1). The seminiferous tubules of all mutant testers had normal spermatogonia, spermatocytes, and Sertoli cells (Fig. 1, B, D, and F). The interstitium between the seminiferous tubules was also normal, with apparently normal Leydig cell number and morphology. Testis histology of Inf4 mutant testers revealed that the spermatogenic abnormality first became obvious at stage VIII, with spermatids in step 8 of spermiogenesis (Fig. 1B). The Inf8 and Inf9 testers began to arrest in stage X, step 10, or slightly later in the program of spermiogenesis (Fig. 1, D and F, respectively).
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We determined whether apoptosis was contributing to the decreased sperm count. Carrier mice from Inf4, Inf8, and Inf9 had very few apoptotic nuclei, all of which were restricted to the spermatogonial layer of the seminiferous tubules (Fig. 2B). In situ cell death detection revealed that in tester mice from all three pedigrees, apoptotic nuclei were localized to the late-differentiating spermatids at the same stage as the spermiogenesis block (Fig. 2, C–F). Transmission electron microscopy of the tubule lumen of the seminiferous tubules of male Inf4 testers also revealed numerous apoptotic nuclei in the tubule lumen (Fig. 2F).
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Sperm Motility and Morphology
A clear distinction was observed in the motility of mutant tester and carrier epididymal sperm from each pedigree (Fig. 3). In particular, Inf4 (0% vs. 84.9% ± 9.5%; P < 0.001), Inf8 (0% vs. 71.2% ± 10.4%; P < 0.001), and Inf9 (0% vs. 83.7% ± 5.2%; P < 0.001) tester males had no forwardly progressive motile sperm (Fig. 3). Because of the severe lack of normal motility, spermatozoa released into the epididymis were analyzed by light microscopy for abnormalities in head and tail morphology. On average, tester animals from Inf4, Inf8, and Inf9 had a significant reduction in normal head and tail morphology compared to carrier mice from each pedigree (Table 2). In particular, sperm from Inf4 tester mice had 55.7% ± 18.5% club-shaped heads, with the most deleterious tail abnormality being stumped short tails (51.0% ± 14.8% in testers vs. 0.3% ± 0.5% in carriers; P < 0.01). Sperm from Inf8 tester mice had 51.4% ± 11.9% club-shaped heads, as well as stumped tails (40.8% ± 11.7% in testers vs. 0.8 + 0.7% in carriers; P < 0.01) and tails with a looped principal piece (27.8% ± 8.9% in testers vs. 1.2% + 1.1% in carriers; P < 0.01). Sperm from Inf9 tester mice had 41.9% ± 12.1% club- shaped heads and severe tail abnormalities, including tails with a looped midpiece (32.1% ± 2.4% in testers vs. 13.1% ± 7.2% in carriers; P < 0.01), a looped principal piece (35.4% ± 2.5% in testers vs. 3.6 ± 1.5% in carriers; P < 0.01), and stumped tails (5.4% ± 1.6% in testers vs. 0 in carriers; P < 0.01).
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Transmission-Electron Microscopy
The lack of motility and presence of severe tail abnormalities visualized by light microscopy prompted a more detailed analysis of tail ultrastructure by TEM. The TEM analysis of Inf9 tester spermatozoan tails in cross-section revealed that four of the nine microtubule doublets were missing from 21% of the tails, with the abnormalities (hemiaxonemes) apparently restricted to the principal piece (Fig. 4, B and C). This same hemiaxoneme arrangement was not observed in cross-sections through the midpiece or endpiece. In comparison, 100% of carrier cross-sections revealed a normal 9+2 microtubule arrangement (Fig. 4A). Analysis of Inf8 spermatozoan tail ultrastructure revealed a related, but distinct, phenotype with an abnormal arrangement of dense fibers (Fig. 4C, arrowhead). We never observed the same hemiaxonemes in Inf8 testers as were observed in Inf9 testers. Ultrastructure of the Inf4 spermatozoan also did not reveal the hemiaxonemes; however, we did observe abnormal manchette assembly (Fig. 4D), which may result in the misshapen sperm heads observed by light microscopy.
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Complementation Analysis
We established a mating scheme to examine whether the infertile mutants formed a noncomplementing group when heterozygous at Inf9 and either Inf4 or Inf8 (Table 3). We determined that Inf9/+;Inf8/+, Inf9/+;Inf4/+, and Inf8/ +;Inf4/+ failed to complement each other when examined for epididymal sperm count and motility (Table 3).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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We determined that the recessive male infertility phenotype was associated with increased apoptosis. Previous studies have revealed that apoptosis associated with disrupted spermatogenesis can be attributed to defects in the apoptotic pathway, such as disruption of the proapoptotic gene Bax [29, 30]. In Bax-deficient mice, an accumulation of premeiotic germ cells in adult mice is associated with a near-complete absence of spermatocytes and mature sperm. In contrast, the increased apoptosis observed in the present study was identified in the late-spermatid stages associated with a defect in spermatid differentiation. Therefore, we hypothesize that the increased apoptosis observed in all three mutant lines results from incorrect sperm differentiation, not from a mutation in an apoptotic pathway.
The late spermatid stage is a critical point in the differentiation of mature sperm. In particular, late-differentiating spermatids undergo significant microtubule rearrangements to form the manchette in the differentiating sperm head and the 9+2 microtubule rearrangement to form the sperm tail. In addition, the nucleus undergoes chromosomal condensation, with histones being replaced by protamines [31] and formation of an acrosomal cap at the tip of the elongating sperm head. We do not know the gene or genes that have been mutated in our ENU mutagenesis strategy; however, all three mutant lines show severe abnormalities in head and tail formation, the most obvious being formation of the 9+2 microtubule arrangement of the tail in two of the alleles, Inf8 and Inf9. The third allele, Inf4, has the most severe defect, resulting in misshapen heads associated with abnormal manchette assembly.
The three mutant lines were identified from three independent pedigrees and, thus, likely represent three independent, ENU-induced mutation events. A breeding scheme was established to produce males that were doubly heterozygous for Inf9 and Inf8, Inf8 and Inf4, or Inf9 and Inf4. These experiments revealed that the three different mutations form a noncomplementing group of alleles. Allelic noncomplementation is identified when compound heterozygous mutations in the same gene have an abnormal homozygous recessive phenotype. Nonallelic noncomplementation is rare, but it is found when heterozygous recessive mutations in different genes act to influence the same phenotypic pathway and, thus, have a phenotype in compound heterozygotes.
The identification of noncomplementing phenotypic groups using ENU mutagenesis was previously reported in a comprehensive study by Rinchik et al. [32]. In their study, 4557 gametes were screened, and allelic noncomplementation was identified at the fit1 (fitness 1), shl (Myo7a), and 17Rn3 loci with the identification of five, seven, and six alleles, respectively. Thus, those authors suggested that the generation of a noncomplementing group containing five to seven alleles after screening 4557 gametes was an average result for a normal locus. In the present study, we have identified a complementation group of three alleles after screening only 184 gametes. In comparison to the study by Rinchik et al., we have an above-average rate of mutation at the locus or loci responsible for the male-factor infertility phenotype on mouse chromosome 11, which could represent 1) loci highly susceptible to amino-acid changes, 2) a large gene, or 3) a clustered gene family. Given that the three mutations are localized to a 24-cM interval on mouse chromosome 11, we predict that the noncomplementing male infertility phenotype is the result of mutations in the same locus or a closely linked genomic cluster of different loci acting in the same pathway.
The most obvious human syndrome with features similar to the Inf4, Inf8, and Inf9 mouse mutants is dysplasia of the fibrous sheath (DFS), or stump tail syndrome [33, 34]. Common characteristics of this syndrome include hyperplasia and marked disorganization of the fibrous sheath as well as axonemal and microtubule doublet distortions [33]. Both complete and incomplete forms of DFS have been observed in the human population, and both are thought to result from an underlying genetic mutation [34]. To our knowledge, no candidate loci for DFS are currently known in humans, and no causal evidence indicates that the two major fibrous sheath proteins, AKAP3 and AKAP4 (located on mouse chromosome 6 and X, respectively), are potential candidates in human patients [35]. Therefore, the interval on mouse chromosome 11/human chromosome 17 between Wnt3 and Trp53 could represent one critical genomic region for DFS.
A mouse mutant with a phenotype that most closely resembles the abnormal sperm morphology described here is the spontaneous mouse mutant azh (abnormal spermatozoon head shape), which arose from mutation in the microtubule attachment protein Hook1 [36–38]. No Hook family members are found on mouse chromosome 11; however, analysis of all known genes within the defined endpoints of the 24-cM balancer reveals four genes with known functions in microtubule and/or cytoskeletal assembly and function. These include Tektin1 (Tekt1), Tektin 3 (Tekt3), Tubulin G (Tubg), and axonemal dynein heavy-chain 2 (Dnahc2). The most likely candidate for this male infertile allelic series is Dnahc2. Dyneins have been shown to play an important role in flagella organization and motility [39]. In particular, axonemal dyneins produce the motive force for ciliary and flagellar beating by inducing sliding between adjacent microtubules within the axoneme. The function of Dnahc2 on mouse chromosome 11 is not known; however, mutation analysis of a related dynein gene, mouse dynein heavy-chain 7 (now called Dnahc1), results in complete male infertility because of abnormalities in sperm tail function [40]. Interestingly, unlike the mutants in the present analysis, the functional abnormality in the Dnahc1 mutants cannot be correlated with morphological changes in axonemal structure. However, another axonemal dynein heavy- chain gene on mouse chromosome 17 (Dnahc8), has also been associated with infertility and abnormal sperm structure [14]. Furthermore, the dynein heavy-chain genes are very large, consistent with the likelihood of isolating an allelic series in the number of families screened. Alternatively, a number of predicted genes in the region of chromosome 11 have no assigned function or gene expression information; thus, these genes may also form potential candidates for the male infertility phenotype.
Taken together, the use of balancer chromosomes with ENU mutagenesis have provided us with a mechanism of ascribing biological function to specific regions of the genome. In the present study, we have identified a noncomplementing male infertility group on mouse chromosome 11. Thus, mouse chromosome 11/human chromosome 17 contains important loci or genomic sequences involved in male infertility and microtubule function and may be one critical region for genetic lesions resulting in DFS.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Correspondence: Monica J. Justice, Baylor College of Medicine, One Baylor Plaza, S413, Houston, TX 77030. FAX: 713 798 1445;mjustice{at}bcm.tmc.edu
Received: 23 June 2003.
First decision: 14 July 2003.
Accepted: 23 December 2003.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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