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Department of Experimental Radiation Oncology (G.S., C.C.Y.W., K.L.P., Z.Z., M.L.M.), The University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030; Department of Physiology (P.P.), University of Turku, 20520 Turku, Finland; and Department of Molecular and Integrative Physiology (T.R.K.), University of Kansas Medical Center, Kansas City, Kansas 66160
Address all correspondence and requests for reprints to: Gunapala Shetty, Department of Experimental Radiation Oncology, The University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030. E-mail: sgunapal{at}mdanderson.org.
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Abstract |
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Introduction |
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The jsd mutation can be a good model for the insight it may provide into the causes and treatments for male sterility caused by other genetic or pathological situations, especially involving spermatogonial arrest. In normal rodents, testosterone (T) and FSH, the two hormones that support spermatogenesis, are involved only in the quantitative maintenance of spermatogonial differentiation and are not critical for the qualitative transformation of type A to intermediate and type B spermatogonia or spermatocytes (5, 6). In contrast, in some pathological situations when the differentiation of spermatogonia is compromised by treatment with radiation (7), chemosterilants (8, 9), heat (10), or aging (11) or by the jsd mutation (12), T and/or FSH appear to be responsible for this block because their suppression by GnRH analogs reinitiates spermatogonial differentiation (13, 14, 15). The involvement of androgen receptor (AR) in this inhibition was indicated by inhibition with other compounds with androgenic activity and the reversal of the inhibition with the AR antagonist flutamide (16). Although FSH was also suppressed by the GnRH-analog treatment, pharmacological studies did not indicate that exogenous recombinant human FSH had an inhibitory role in spermatogonial differentiation in jsd mutant mice (15). However, in irradiated LBNF1 rats, another model of spermatogonial block, we have observed inhibition by exogenous FSH and endogenous FSH levels (17).
The goal of the present study was to test the AR-mediated inhibition of spermatogonial differentiation by T and any possible inhibitory function of endogenous FSH in jsd mutant mice through genetic approaches. To eliminate the AR, we used testicular feminized (Tfm) mice (18), which have a mutation within the coding region of the Ar gene that leads to an inactive receptor (19). Tfm males have small abnormal testes, located on either side of the bladder in the abdomen, with tubules containing Sertoli cells, spermatogonia, and some spermatocytes, which are lost during meiotic prophase (20). To specifically eliminate FSH action, we used fshb–/– mice (21). Adult male fshb–/– mice have scrotal testes, with qualitatively normal spermatogenesis, and are fertile, although testis weights and sperm production are 40 and 25%, respectively, of that of normal littermates. The reduced testes weight and sperm production in fshb–/– mice result from a drop in the number of Sertoli cells and further decline in germ cell numbers, producing a spermatogonia/Sertoli cell ratio of 0.36 compared with a value of 0.47 in wild-type mice (22). For this study, Tfm and fshb–/– mice were independently crossed to jsd mice, and spermatogonial differentiation was assessed in the double mutants with or without additional treatments.
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Materials and Methods |
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Tfm/+ heterozygous females on C57BL6/J (B6) background (obtained from Jackson Laboratory, Bar Harbor, ME) were first backcrossed with C3H males, and female Tfm/+ offspring, which had the C3H haplotype flanking the jsd locus, were selected. This was necessary because the jsd mutant allele was initially identified by PCR amplification of B6- and C3H-specific polymorphic microsatellite loci tightly linked to the jsd locus (14). These Tfm/+ female mice were crossed to jsd/+ males, which were on a B6-C3H mixed background developed by crossing C3H mice with mutant jsd-carrying B6 mice. The resulting jsd/+,Tfm/+ females were again crossed to jsd/+ males to obtain jsd/jsd,Tfm/Y males (note Ar is present on the X chromosome). For comparison (jsd or +)/+,Tfm/Y, and jsd/jsd,+Y littermates were used. There were no testicular phenotypic differences between jsd/+ and wild-type mice as indicated by the lack of significant differences in testes weights of these mice and the fathering of normal numbers of pups by jsd/+ mice.
Similarly, male fshb–/– mice (fshbm1/fshbm1) on a 129-B6 background (21) were initially crossed with C3H females, and offspring with the C3H haplotype at the jsd locus were selected. The fshb–/– males, produced by intercrossing these mice, were bred to jsd/jsd females on C3H-B6 background. The double heterozygotes produced were intercrossed to obtain jsd/jsd,fshb–/– males and (jsd or +)/+,fshb–/– and jsd/jsd,fshb+/– littermates.
Screening the mice for jsd, Tfm, and fshb-null mutations
Initially, jsd mice were genotyped by PCR analysis of crude tail DNA from animals at 3–5 wk of age, using loci D1Mit 415 and D1Mit 181, which encode microsatellite DNA sequences close to the jsd locus (23). Subsequently, after the jsd gene was identified, the genotyping was done using purified DNA using the primers described and testing the PCR product for sensitivity to HphI digestion (3).
The Tfm mice were genotyped by PCR analysis as described earlier (24) with a minor modification of a change in the buffer used to prepare the PCR mixture (23). Fshb mutant mice were also screened by PCR following previous procedures (25, 26) except that the forward and reverse primers used for hprt (hypoxanthine-guanine phosphoribosyltransferase) were 5'-CCC CAC GAA GTG TTG GAT ATA A-3' and 5'-AGG CTC ATA GGG CAA ATA AAC A-3'. The presence of hprt indicates the presence of the fshb-null allele, because this allele was generated by replacing exons 1 and 2 and most of exon 3 by the PGK-hprt expression cassette (21).
Experimental design
The age dependence of spermatogenesis was assessed in jsd mutant mice also carrying the Tfm mutation. Mice were killed at the ages of wk 12 or 20 to analyze spermatogonial differentiation and hormone levels. Next, the effect of orchiopexy in these jsd/jsd,Tfm/Y mice was assessed. Orchiopexy of the right testis was performed at the age of 6–8 wk, and the mice were killed 8 wk later for the analysis of spermatogonial differentiation and serum T levels.
With jsd mice also carrying an fshb-null mutation, the age dependence of the decline in spermatogenesis was assessed first. The mice were killed at 5, 8, and 12 wk of age to analyze spermatogonial differentiation. Hormone analysis was also done in these mice killed at wk 12. Next, the effects of hormone suppression in these jsd/jsd,fshb–/– mice were determined. The mice were treated with a GnRH and an AR antagonist starting at the age of 8 wk and killed at 12 wk to analyze spermatogonial differentiation and hormone levels.
In all these experiments, single-mutant littermates and wild-type mice served as controls.
Histological analysis
In all experiments, testes were weighed. In untreated and hormone-treated mice, the left testis was fixed in Bouin’s fluid and embedded in paraffin or methacrylate, and sections were stained in hematoxylin or periodic acid Schiff (PAS)-hematoxylin; the right testis was used for intratesticular T (ITT) measurements. All seminiferous tubules (approximately 100–150) in the testis section were categorized as either differentiating (containing germ cells at the stage of B spermatogonia or beyond) or not, and the tubule differentiation index (TDI) was calculated as the percentage of differentiating tubules. The numbers of A spermatogonia/100 Sertoli cells, apoptotic indices, and mitotic indices were measured as described earlier (27) in methacrylate sections of testes from 12-wk-old jsd/jsd and jsd/jsd,fshb–/– mice. The morphological criterion used for apoptosis was the appearance of dense chromatin bodies distributed in the cell nucleus.
Hormone treatment
To suppress T levels and androgen action, we gave jsd/jsd,fshb–/– mice a combination of GnRH antagonist and flutamide for 4 wk starting at 8 wk of age. To determine the dose of the GnRH antagonist acyline (Contraceptive Development Branch of National Institute of Child Health and Human Development, North Bethesda, MD) that was required to suppress ITT levels, we performed preliminary experiments using normal adult C3H mice, giving different doses of acyline in various concentrations. The acyline was dissolved in sterile water, and the volume of acyline solution injected sc at each site was kept constant: 0.5 ml/100 g body weight. Single doses of 0.6 mg/kg and 2 mg/kg did not significantly suppress ITT levels (data not shown). When single doses of 5, 10, and 20 mg/kg body weight were given in concentrations of 1, 2, and 4 mg/ml, respectively, the suppression of ITT at 1 wk showed an inverse relationship with the doses used (Fig. 1A
). Thus, the initiation of ITT suppression was delayed with increasing doses of acyline, although the suppression was maintained for longer periods in mice treated with higher doses. For example, the 10-mg dose produced about 2 wk of suppression, but the ITT levels were still suppressed to 5% of the control levels at 3 wk in the 20-mg/kg group. These results support the observation that at concentrations of 2 mg/ml or more, acyline bioavailability is delayed, presumably because of the formation of a gel (Hild, S., personal communication). Hence, for the efficient suppression of ITT levels, we gave two sc injections each of 10 mg/kg at separate sites in a concentration of 2 mg/ml. This dose suppressed the ITT to 9% of the control level within a week and further reduced ITT to 2% of the control levels at 2 wk. To maintain suppression of ITT, a second single injection (10 mg/kg) of acyline was given 2 wk after the first set of injections; this second dose sustained suppression for at least another 4 wk.
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). Based on these results, the jsd/jsd,fshb–/– mice were treated with two injections of acyline (each 10 mg/kg) at 8 wk of age and simultaneously implanted with two 2-cm flutamide implants, which remained in place until the mice were killed at 12 wk of age. An additional acyline injection of 10 mg/kg was given at 10 wk of age to maintain suppression of T production.
Surgical manipulation of mice
Testes of jsd/jsd,Tfm/Y, jsd/+,Tfm/Y, and jsd/jsd,+/Y mice were translocated to a sc pouch at the age of 6–8 wk. Because Tfm/Y mice have abdominal testes and lack a scrotum, a scrotal pouch could not be created. We made an incision in the lower abdomen, and the right testis along with the fat pad was pulled out through a hole made in the peritoneal wall and secured in a sc position by suturing either the fat pad or the tunica of the testis to the outer peritoneal wall. Care was taken not to damage the blood vessels during this procedure, and mice with fibrosed atrophic testis observed at the time of analyses were excluded from the study. In jsd/jsd,+/Y mice, the right testis, which was initially in the scrotum, was pulled up to the lower abdominal region, followed by the above procedure, to place the testis in an anatomical position similar to that of the surgically manipulated Tfm/Y mice. The mice were killed 8 wk after surgery for the analyses of the left testis, which was in its original position, and the right sc testis. The observation of differentiated spermatogenic cells in most of the surgically manipulated testes and the appearance of fibrosis and atrophy in only 15% supports our ability to reliably translocate the testis without damage to blood vessels.
Hormone measurements
At the time of killing, the mice were anesthetized with sodium pentobarbital, and blood was collected from the axillary vein for analysis of T and/or gonadotropins. The serum was separated and stored at –20 C. The entire right testis or a portion of each testis was weighed, immediately frozen in liquid nitrogen, and stored at –80 C. The testis was homogenized, debris was removed by centrifugation, and the supernatant was used for the assay of ITT. Serum and ITT were measured using the DSL-4000 (Diagnostic Systems Laboratories, Webster, TX) coated-tube RIA kit as described earlier (13). The lower limit of detection for T by this assay was 0.041 ng/ml. Serum levels of FSH were measured using immunofluorometric assay (Delfia; Wallac Finland Oy, Turku, Finland) as previously described (29).
Statistical analysis
The data are represented as the arithmetic mean ± SEM for TDI, testis weight, and serum FSH and as the averages and SEM calculated on log-transformed data for ITT and serum T. The differences between the groups were first analyzed by one-way ANOVA. If the differences were significant (P < 0.05), Dunnett’s post hoc test for multiple comparisons was performed to determine the significance of differences between wild-type or double mutants and the jsd/jsd mutant group. When only two groups were compared, a t test was performed instead to analyze the significance of the difference. In comparisons of the values for the left and right testis of the same animal, a paired-samples t test was performed. A computer-assisted statistics program (SPSS, Inc., Chicago, IL) was used.
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, C and D, and Table 1). Age-matched jsd/jsd,+/Y mice on the C3H-B6 mixed genetic background showed differentiation in only 14% of the tubules; 11% of the tubules contained spermatocytes and the rest earlier stages (Fig. 2, A and B). In double jsd/jsd,Tfm/Y mutants, the testis appeared similar to that of Tfm/Y mice, with 94% of the tubules showing differentiation; 89% of them contained spermatocytes and 5% only B spermatogonia (Fig. 2, E and F). Germ cell differentiation was also maintained in 20-wk-old jsd/jsd,Tfm/Y mice, which showed a TDI of 98%, compared with 2 and 96% in jsd and Tfm single mutants, respectively. Thus, spermatogonial differentiation was restored in adult jsd mutant mice with defective AR.
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). Second, the testes in jsd/jsd,Tfm/Y mice are in a cryptorchid position, and cryptorchidism was recently shown to restore spermatogonial differentiation in jsd mutant mice without a concomitant reduction in hormone levels (23).
To eliminate the low levels of T as a possible factor in the release of inhibition of spermatogonial differentiation in jsd/jsd,Tfm/Y mice, we treated these mice with 2-cm T capsules from 4–12 wk of age. The sc delivery of T elevated serum T levels from 0.5 to 21 ng/ml in jsd/jsd,Tfm/Y mice resulting in an ITT concentration of 14 ng/g testis (Table 1). Although the ITT concentrations were not significantly increased, they were still within the range of the serum T concentrations. We had previously shown that a 4-wk treatment regimen of only 0.5-cm T capsules produced ITT concentrations of 19 ng/g testis in GnRH antagonist-treated jsd/jsd mice and reduced the TDI from 73 to 7% through an AR-mediated process (14). Similar ITT levels with high serum T concentrations did not alter the TDI in these jsd/jsd,Tfm/Y mice, demonstrating that non-AR-mediated inhibition of spermatogonial differentiation by T did not occur in these mice.
In an attempt to eliminate the elevation of temperature as a factor, we placed the testis of jsd/jsd,Tfm/Y mice in a sc pouch for 8 wk by performing unilateral orchiopexy. In these mice, the TDI was not altered in the sc placed testis (97%) compared with the intact contralateral abdominal testis (99%) (Table 2). However, after a similar orchiopexy, even the testes from jsd/jsd,+/Y mice showed differentiation in almost all the tubules, suggesting that temperature in such a sc pouch was still above scrotal temperature. We had previously observed that in 12-wk-old jsd/jsd mice, testes placed in the abdomen for a period of 8 wk weighed 45 ± 5 mg (23), whereas scrotal testis weighed 20 ± 1 mg. In the present study, the testis of jsd/jsd mice after orchiopexy weighed 41 ± 4 mg, which was similar to the abdominal testis.
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). The testis weights were significantly reduced in jsd/jsd,fshb–/– mice compared with jsd single mutants, which was likely a direct effect of FSH deficiency on somatic cell development. None of the tubules showed spermatogenic cell differentiation in jsd/jsd,fshb–/– mice at the age of 12 wk (Fig. 3, E and F), which was similar to the value of 1% in jsd single-mutant mice on this C3H-B6–129 mixed genetic background (Fig. 3, C and D), suggesting FSH did not contribute to the inhibitory effect on spermatogonial development in jsd mutants (Table 3). Nevertheless, to confirm the conclusion, it was necessary to show that sufficient numbers of type A spermatogonia were present in these smaller testes and that they could differentiate under appropriate conditions.
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We next examined the numbers of spermatogonia and their kinetics in adult jsd/jsd,fshb–/– mutants at 12 wk of age. Although there seemed to be fewer spermatogonia in the double mutants (3.9 ± 1.0 per 100 Sertoli cells) than in jsd/jsd single mutants (9.3 ± 3.7 per 100 Sertoli cells), the difference was not significant. The proliferative activity of spermatogonia in jsd/jsd,fshb–/– testes was unaltered as indicated by their high mitotic index (35.5 ± 4.2%), which was similar to that in jsd single mutants (25.5 ± 2.4%). They also underwent apoptosis instead of differentiating, producing an apoptotic index of 6.2 ± 1.9%, just as was the case in jsd single mutants (apoptotic index of 6.5 ± 1.9%).
The jsd/jsd,fshb–/– mice had very high ITT concentrations mainly because of the reduced size of the testis (Table 3). The ability of the spermatogonia in these mice to differentiate when such levels of T and its action were suppressed was examined by giving a combination of GnRH antagonist and flutamide (Table 3). Treatment significantly suppressed the ITT and serum T concentrations in these mice by more than 96% and restored spermatogonial differentiation in 63% of tubules, which was not significantly different from the value in jsd/jsd mice (TDI = 76%). These results show that spermatogonia in jsd/jsd,fshb–/– mice were capable of differentiation.
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Discussion |
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). The percentages of tubules showing differentiation in GnRH-antagonist- and flutamide-treated jsd/jsd and jsd/jsd,fshb–/– mice were similar, although the GnRH-antagonist-treated jsd single-mutant mice had an appreciable amount of serum FSH, about 3 ng/ml, confirming that FSH neither inhibits nor stimulates spermatogonial differentiation in jsd mutant mice. FSH’s ability to inhibit differentiation in the irradiated rat (17) but not in the jsd mutant mouse could be because of the involvement of different mechanisms in the block of spermatogonial differentiation. In rat, it may be mediated by Sertoli cells and in mouse by other androgen-responsive cells in the testis. The demonstration of the loss of inhibition of spermatogonial differentiation in jsd mutant mice by elimination of AR was primarily a result of failure of AR-mediated testicular descent that is ordinarily caused by the contraction of the gubernaculum. Temperature elevation in the cryptorchid Tfm mutant testis largely bypasses the T-induced block in spermatogonial differentiation (23). We were unable to reduce the testicular temperature sufficiently to be able to critically evaluate whether T, acting through the AR in testicular cells, would have a direct inhibitory effect on spermatogonial differentiation in jsd/jsd,Tfm/Y mice. Even when the testes were placed in a sc pouch, the temperature was still higher than the scrotal temperature and the elevation was sufficient to bypass the T-induced block, as indicated by the restoration of spermatogonial differentiation in jsd/jsd mice.
In other studies, orchiopexy at least partially restored spermatogenesis, such as in mice with genetic mutations in Insl3 (31) and its receptor, Lgr8 (32). Tubules with complete spermatogenesis were observed in Insl3-mutant mice when orchiopexy was performed by placing the testis in a scrotal pouch (31), whereas maturational arrest was observed when the testis was placed in a sc pouch. The other study also involved the translocation of the abdominal testis of Lgr8-mutant mice to a superficial skin pouch in the region of the scrotum and resulted in tubules with complete spermatogenesis (32). However, in our study, such a scrotal pouch could not be created in Tfm male mice because they do not develop a scrotum. Hence we may not have been able to reduce the testicular temperature as much as when the testes were transplanted to the scrotum. In a different approach, it has been shown that testicular tissue grafts from prepubertal animals when transplanted sc into nude mice can support qualitatively normal spermatogenesis (33). Although the lack of fur may lower the temperature, this observation suggests that greater reduction in temperature is required to produce the block in spermatogonial differentiation in jsd mutant mice than to maintain spermatocyte and spermatid differentiation. A conditional knockout of AR will be required to produce mice with normally descended testis but no AR in somatic cells of the testis (34) to test whether the receptor-mediated inhibition of spermatogonial differentiation by T involves action on testicular somatic cells and to determine the specific cell type involved.
The present data provide additional information regarding the effect of genetic background on the severity of jsd phenotype. For example, at 12 wk of age, jsd mutants that were on a B6-C3H background (jsd single-mutant littermates of jsd-Tfm double mutants) had a TDI of 14%, whereas mice on a B6–129-C3H background (jsd single-mutant littermates of jsd-fshb double mutants) had a TDI of 1%. Additional studies are in progress to test whether genetic background affects the severity of the jsd phenotype in inbred mice (Bolden-Tiller, O. U., H. Chiarini-Garcia, and M. L. Meistrich, unpublished data).
Our data indicate that FSH does not contribute to the inhibition of spermatogonial differentiation in jsd mice. We have also shown that elevation of testicular temperatures relieves the inhibition of spermatogonial differentiation.
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Acknowledgments |
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Footnotes |
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The authors have nothing to declare.
First Published Online April 20, 2006
Abbreviations: AR, Androgen receptor; ITT, intratesticular testosterone; PAS, periodic acid Schiff; T, testosterone; TDI, tubule differentiation index.
Received February 8, 2006.
Accepted for publication April 10, 2006.
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and androgen receptor in the regulation of neuronal nitric oxide synthase. J Comp Neurol 453:336–344[CrossRef][Medline]This article has been cited by other articles:
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  G. Shetty, S. H. Shao, and C. C. Y. Weng p53-Dependent Apoptosis in the Inhibition of Spermatogonial Differentiation in Juvenile Spermatogonial Depletion (Utp14bjsd) Mice Endocrinology, June 1, 2008; 149(6): 2773 - 2781. [Abstract] [Full Text] [PDF]
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 O. U. Bolden-Tiller, H. Chiarini-Garcia, C. Poirier, D. Alves-Freitas, C. C. Weng, G. Shetty, and M. L. Meistrich Genetic Factors Contributing to Defective Spermatogonial Differentiation in Juvenile Spermatogonial Depletion (Utp14bjsd) Mice Biol Reprod, August 1, 2007; 77(2): 237 - 246. [Abstract] [Full Text] [PDF]
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), 10 mg/kg (
), or 20 mg/kg (
) body weight on d 0 or two injections of 10 mg/kg body weight at different sites on d 0, followed by no additional injections (
) or followed by a single injection of 10 mg/kg body weight on d 14 (
) (n = 5 mice per point); 
, value for untreated mice. Significance of difference from the control untreated mice is P < 0.001 for all the points below the dotted line. B, Reduction of sperm head counts in mice given two simultaneous injections of acyline at 10 mg/kg each with additional flutamide treatment, given via sc implanted capsules of indicated lengths (
) starting on d 0. Sperm heads were counted 2 wk later along with acyline-only-treated (
