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Original Studies |
Prince Henry’s Institute of Medical Research (L.O., G.B., D.M.R., R.M.), Clayton, Victoria 3168, Australia; Population Center for Research in Reproduction (A.N., Y.-Q.G., W.J.B.), Department of Medicine, University of Washington, Seattle, Washington 98195; and Department of Anatomy and Cell Biology (N.G.W.), Monash University, Clayton, Victoria 3168, Australia
Address all correspondence and requests for reprints to: Liza O’Donnell, Ph.D., Prince Henry’s Institute of Medical Research, P.O. Box 5152, Clayton, Victoria 3168, Australia. E-mail: liza.odonnell{at}med.monash.edu.au
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Abstract |
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Introduction |
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Although many studies have investigated the effects of hormone suppression on spermatogenesis in the rat (5, 6, 7), fewer studies have investigated the effects of FSH and LH/testicular T suppression on germ cell development in monkeys and man. In monkeys rendered gonadotropin-deficient by hypophysectomy or GnRH antagonist treatment, spermatogonia are the most sensitive germ cell type (8, 9, 10, 11); however, which spermatogonial subtypes are regulated by hormones is controversial (9, 11, 12, 13, 14). A recent study in men receiving weekly injections of T enanthate suggested that the inhibition of spermatogonial development was also the primary lesion in response to gonadotropin withdrawal and further suggested that the failure of sperm release may also be important (15). In accordance with this latter observation, our recent study in rats showed that spermiation failure contributes to the suppression of spermatogenesis after hormone withdrawal (16).
Potential T-based contraceptive regimens in monkeys and humans clearly inhibit various phases of germ cell development; however, the relationship between these changes and the extent of suppression of sperm output is unclear. Our recent results in monkeys given T implants indicated that animals who achieve stable azoospermia in response to 20 weeks of T administration have significantly lower levels of circulating FSH, compared with those animals who did not achieve consistent azoospermia, whereas no differences were seen in LH or T (serum or testicular) levels (17).
The purpose of this study is to describe the time course of the effects of exogenous T administration on germ cell development in macaque monkeys and to relate the degree of suppression of germ cell development to sperm output. The optical disector stereological technique was used for the unbiased determination of Sertoli cell and germ cell number in testicular biopsies taken before, during, and after T administration. The data show that reductions in type A pale (Ap) and B spermatogonia are the key lesions in spermatogenesis and that failure of spermiation is also important.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Nine normal adult male crab-eating macaque monkeys, Macaca fascicularis (age, 5–12 yr; BW, 5–9 kg) were housed in the Regional Primate Research Center at the University of Washington, under defined environmental conditions. All procedures were performed in accordance with the Endocrine Society Guidelines for the Care and Use of Experimental Animals (18). All procedures were carried out under Ketamine (Fort Dodge, IA) HCl (10 mg/kg, im) anesthesia. Valium (1 mg/kg; im, Elkins-Sinn, Inc., Cherry Hill, NJ) was given, before Ketamine HCl, to all monkeys, for the electroejaculation procedure.
The study involved a pretreatment (control) phase (4 weeks), treatment phase (20 weeks) terminating with the removal of the implants, and recovery phase (16 weeks). T was administered via sterilized sc SILASTIC implants (5.5-cm long; inside diameter, 0.33 cm; outside diameter, 0.46 cm ) for 20 weeks. Two implants were inserted sc in the midscapular region of each animal.
Testicular biopsies
Open testicular biopsies were performed alternatively on the right and left testis, in aseptic conditions, under ketamine/xylazine anesthesia. All animals were biopsied 4 weeks before the treatment (control) and then at weeks 2, 14, and 20 during the treatment and at 12 weeks post treatment. Approximately 300–500 mg tissue was taken per biopsy and was divided in half; half was frozen in liquid nitrogen for the measurement of testicular androgens (Narula et al., submitted). The other half was immersion-fixed in Bouin’s fluid for approximately 5 h, then stored in 70% ethanol. Biopsies were observed under a dissecting microscope and cut along a plane lying at right angles to the dominant tubule orientation, to maximize the number of circular tubule profiles.
Testicular function and hormone data
Body weights, testis volumes, and sperm counts were assessed every 2 weeks during control, treatment, and recovery periods. Testis volumes were measured using an orchidometer. Seminal fluid was collected after rectal probe electroejaculation, and total sperms counts (millions/ejaculate) were determined using a hemocytometer. Blood samples were collected from the femoral vein every 2 weeks during the study (Narula et al., submitted). Bioassays were used to measure the bioactivity of LH and FSH as detailed elsewhere (Narula et al., submitted). The detection limits of the assays were as follows: bioLH, 2.4 IU/L; bioFSH, 0.08 IU/L. Testicular T was measured using previously published methods (19) as detailed elsewhere (Narula et al., submitted).
For the purposes of the current study, hormone and semen data are included only from the time points where biopsies were taken to compare with stereological data. More detailed data on the endocrine parameters and sperm output every 2 weeks across the entire study are reported elsewhere (Narula et al., submitted).
Tissue processing
Bouin’s fixed biopsy tissue was dehydrated through a series of ethanol and butanol steps, then processed and embedded in hydroxyethylmethacrylate resin (Technovit 7100, Kulzer and Co., Friedrichsdorf, Germany), according to the manufacturer’s instructions. Thick (25-µm) sections were cut using a Supercut microtome (Reichert Jung 2050, Nussloch, Germany) and stained with PAS (periodic acid-Schiff’s reagent) and counterstained with hematoxylin.
Cell identification and groupings
Each seminiferous tubule was staged into one of the following stage groupings based on the cellular associations present in stages I-VI, VII, VIII-XI, and XII (see (20), and the cells were counted in the following groups: Sertoli cells, type A dark (Ad) spermatogonia (present in all stages); type Ap spermatogonia (present in all stages); type B (B) spermatogonia (present in stages X-XII and I-VI); preleptotene + leptotene + zygotene (Pl-Z) primary spermatocytes (present in stages VII-XI); pachytene primary spermatocytes (PS) (present in all stages); round spermatids (rSt 1–7) (stages I-VII); elongating spermatids (eSt 8–12) in steps 8–12 (stages VIII-XII); and elongated spermatids (eSt 13–14) in steps 13–14 (stages I-VI).
Sertoli cells and germ cells, in 25-µm thick PAS-stained methacrylate
sections, were classified on the basis of their nuclear morphology, as
previously described by Zhengwei (20), based on the
criteria of Clermont (21, 22). Spermatogonia were
subdivided into type Ad and type Ap spermatogonia, and type B
spermatogonia as follows (see Fig. 1
for
morphology of the various subclasses of spermatogonia):
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Ap spermatogonia. Ap spermatogonia had nuclei that were
generally larger than Ad spermatogonia and were paler in color. The
nuclei were almost always round and showed one or two clumps of
chromatin associated with the nuclear membrane. The cytoplasm did not
show PAS positive material (Fig. 1).
B spermatogonia. B spermatogonia had nuclei that were
rounded, darker in color, with peripherally located clumps of chromatin
and a central nucleolus. The nuclei were situated further away from the
basement membrane than Ad spermatogonia (Fig. 1).
Sertoli cells were clearly distinguishable from spermatogonia by their
irregularly shaped pale-colored nucleus and the presence of a single
prominent nucleolus (see Fig. 1). Criteria for classification of the
later germ cell types have been described in detail in our previous
study (20).
Stereological analysis
The optical disector method was used to determine the numbers of germ cells and Sertoli cells in each biopsy as previously described (20, 23). This method is an unbiased and efficient technique for estimating cell number, including nonspherical objects such as elongated spermatids, and assumes only that cell number equates to nuclear number. The 25-µm methacrylate sections were optically sectioned by means of a high numerical aperture lens (100x objective lens, numerical aperture 1.4) on an Olympus Corp. (Tokyo, Japan) BX-50 microscope equipped with a microcater (Heideinhain D83301; Traunreut, Germany) attached to the microscope stage to monitor the depth scanned. The image was captured by a Pulinix TMC-6 video camera coupled to a Pentium personal computer using a Screen machine II fast multimedia video adapter (FAST, Hamburg, Germany). A software package, DH CASTGRID version 1.10 (Olympus Corp., Munich, Germany) was used to superimpose a set of unbiased counting frames on the video image. Fields to be counted were selected by a systematic uniform random sampling scheme, with the use of a motorized stage (Multicontrol 2000, ITK, Lahnau, Germany). In each field, 4 counting frames (area of each frame, 460.5 µm2) were generated; cells that appeared infrequently (e.g. spermatogonia, preleptonene-to-zygotene spermatocytes, and Sertoli cells) were counted in each of the 4 frames (total area, 1842 µm2), whereas cells appearing more frequently (such as spermatids) were counted in one frame. In most cases, more than 100 cells were counted per cell grouping per biopsy. However, in some treated biopsies where spermatogenic suppression had occurred, approximately 40 cells were assessed in some germ cell categories.
During counting of germ cell and Sertoli cell nuclei, the position of a fixed point, at -3 µm in the z-axis, was determined in each field to assess the volume fraction of interstitium, seminiferous tubule, and stage of seminiferous tubule. The volume fraction of each stage grouping (I-VI, VII, VIII-XI, and XII) was calculated for each biopsy as an approximation of the relative frequency of each stage. In all control and T-treated testis biopsies counted, the relative stage frequencies obtained were compared with the published relative stage frequency, as determined by Fouquet and Dadoune in Macaca fascicularis (24) to ensure constant recognition of stages (and thus, accurate cell identification). Relative stage frequencies were also used to assess the within-biopsy and interanimal variation. The within-biopsy variation was shown to be 5–14% (n = 9) for each stage grouping, and the interanimal variation was shown to be 9–19% (n = 9). In all treated biopsies, the relative stage frequencies for each stage grouping were similar to the published relative stage frequencies (24) and to the mean values obtained from nine control animals.
The numerical density (Nv) of each cell type was calculated as previously described (23). The Nv of each germ cell type was divided by the Nv of Sertoli cells to express the number of germ cells per Sertoli cell (N/SC). The use of the latter in evaluating the effects of hormones on spermatogenesis relies on the assumption that there are no changes in the number of Sertoli cells per testis. The validity of this assumption has been demonstrated previously in primates (Macaca fascicularis) receiving a GnRH antagonist (11). To compare germ cell types at the one time point, the percent control value was calculated by dividing N/SC of each germ cell type, during the treatment and recovery period, by the N/SC of the control biopsy. Conversion ratios (CR) were also used to assess differences in germ cell development between control and treated biopsies at each time point and were calculated by dividing the N/SC of one germ cell type by the N/SC of the preceding germ cell type in the cycle of spermatogenesis. The CR for each transition was then compared with the CR obtained for the control biopsies. CR were assessed in the 14- and 20-week testicular biopsies where stable spermatogenic suppression had been achieved. It should be noted that the CR were not adjusted for the relative stage duration; and therefore, these ratios cannot be used to assess the efficiency of spermatogenesis as in other studies (20). Rather, they were used to assess lesions to germ cell development by investigating significant differences between control and treated biopsies.
Assessment of spermiation failure
The release of mature step 14 spermatids via spermiation occurs at the end of stage VI/early stage VII; and thus, mature step 14 spermatid heads located in the basal portion of the epithelium from stage VII and beyond represent retained spermatids that do not undergo spermiation. The number of retained spermatids in stages VII–XII were counted, as per other germ cells, and expressed per Sertoli cell.
To compare the number of spermatids retained within the epithelium, with the number of normal spermatids available for spermiation, the number of retained spermatids in stages VII-XII and steps 13–14 spermatids in stages I-VI (N/SC) were corrected for the different duration of stages in which they are present, and were thus multiplied by the relative stage frequency for that stage grouping. The number of retained spermatids per Sertoli cell in stages VII-XII was then divided by the number of steps-13–14 spermatids (in stages I-VI) per Sertoli cell and multiplied by 100, as an estimation of the percentage failing to spermiate. It should be noted that the comparison of retained spermatids to steps 13–14 spermatids is only an approximation of the percentage of spermatids failing to spermiate; because retained spermatids will undergo phagocytosis and disappear over the 136-h period from stage VII to XII, and thus the number of retained spermatids counted will be an underestimation of the actual number retained. However, because of the severity of spermatogenic suppression in most animals, it was not possible to subdivide between these stages; and thus, all retained spermatid data were pooled.
Spermiation failure was only assessed in animals that had spermiogenic cells present in the biopsy during the T-administration period. Because three animals had severe spermatogenic arrest and no elongating or elongated spermatids present in the testis at either the 14- or 20-week treatment periods, these animals were excluded from this analysis.
Statistics
For comparison of one endpoint across the study period, data were log-transformed and tested for homogeneity of variance before repeated-measures one-way ANOVA. When a P value of less than 0.05 was obtained by ANOVA, post hoc analysis was performed using Dunnett’s test for differences compared with the control group, and Newman-Keuls post hoc comparison was performed for other differences between the various time points. When comparing between different germ cell populations, expressed as percent control, the loss of germ cells at different points in the spermatogenic cycle was associated with significant heterogeneity in variances; and thus, nonparametric analyses for correlated samples (Friedman ANOVA and Wilcoxen signed-rank test) were used. When comparing data between groups of animals achieving either consistent azoospermia (azoo group) and those that did not (nonazoo group), significant heterogeneity of variance was observed; and thus, the nonparametric Mann-Whitney U test was used. Differences at the level of P < 0.05 were considered significant. Data are expressed as mean ± SEM (n = 9 for all groups except the 14-week treatment group, where n = 8).
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Results |
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The effect of T administration on the number of each germ cell
type (N/SC), in comparison with control values for all nine animals, is
shown in Table 1. After 2 weeks of T
administration, there were no significant changes in germ cell
populations as grouped in the categories shown in Table 1. After 14 and
20 weeks of T administration, significant decreases were seen in all
germ cell populations from B spermatogonia through to steps 13–14
elongated spermatids. The number of A spermatogonia (Ad + Ap
spermatogonia) was significantly below control only at the 20-week time
point. All germ cell types recovered within 12 weeks after removal of T
implants.
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). The total number
of type A spermatogonia remained between 73–99% of control, across
the treatment period. After 14 and 20 weeks of T administration, the
number of type B spermatogonia was suppressed to 20–38% of control
values. All later germ cell types were suppressed to a similar
degree.
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Effect of T administration on sperm counts and hormone levels
T administration caused significant decreases in sperm counts,
intratesticular T concentrations, and the levels of circulating
bioactive FSH and LH (Table 1). All parameters returned to control
levels within 12 weeks after the removal of T implants. All animals
achieved maximal suppression of sperm counts between 10–16 weeks of
treatment (Narula et al., submitted).
Effect of T administration on spermatogonial subtypes
In the monkey, the type A spermatogonia population is comprised
broadly of two morphologically distinct classes of spermatogonia. Ad
spermatogonia (see Fig. 1) are considered to be the nonproliferative
reserve spermatogonial population (21, 22, 24), whereas
the Ap spermatogonia (see Fig. 1) are considered to provide the
differentiating cells committed to spermatogenesis (21, 22, 24, 25). The effects of T administration on type Ad, Ap, and B
spermatogonia were considered (Fig. 3).
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, Fig. 3), changes were seen when the type A spermatogonial
subtypes were considered separately. As shown in Fig. 3, type Ap
spermatogonia were suppressed to 45% of control (P <
0.05), and Ad spermatogonia increased to 165% of control
(P < 0.05). Despite these changes, no change was seen
in B spermatogonia at this time.
After 14 weeks of T administration, Ap spermatogonia remained
suppressed, whereas Ad spermatogonia had declined to control levels
(Fig. 3). Type B spermatogonia were now suppressed to an extent similar
to that of the Ap spermatogonia (38% and 32% of control for Ap and B,
respectively). After 20 weeks of T administration, the number of Ap
spermatogonia had risen slightly, whereas the number of Ad
spermatogonia decreased further (compared with 14 weeks). When the
total A spermatogonial number was considered, there was a significant
decrease at 20 weeks. Type B spermatogonia remained suppressed after 20
weeks of T administration. When the CR between the spermatogonial
subtypes were analyzed, the only significant decrease was seen in the
transition of Ap to B spermatogonia after 20 weeks (not shown).
Twelve weeks after removal of the T implants, the total number of A
spermatogonia returned to control values (Fig. 3). Ad spermatogonia
were significantly elevated above control, whereas Ap spermatogonia
were significantly below control values.
Failure of spermiation during T administration
There was a significant linear relationship between sperm count and the number of steps 13–14 elongated spermatids per Sertoli cell, after both 14 (r = 0.93, P < 0.001, n = 8) and 20 weeks (r = 0.81, P < 0.01, n = 9) of T administration. However, in some animals, the sperm count did not reflect the number of mature spermatids (steps 13–14) in the testis. For example, animal no. 97114 was azoospermic at 18 and 20 weeks but had 35% the control number of steps 13–14 elongated spermatids in the testis, suggesting a failure of spermiation. Therefore, the failure of spermiation was assessed during T administration. Because three animals had severe spermatogenic suppression after long-term T administration, and no elongating or elongated spermatids were present in their testes, these animals were excluded from the analysis of spermiation.
In control testicular biopsies, few mature retained spermatids were
seen in the basal portion of the epithelium (0.023 ± 0.016 N/SC,
n = 6), and similar numbers were seen after 2 weeks of T
administration (0.046 ± 0.016, n = 6). After 14 and 20 weeks
of T administration, the numbers of retained spermatids per Sertoli
cell increased to 0.071 ± 0.018 (n = 5) and 0.235 ±
0.179 (n = 6), respectively; however, there was marked
between-animal variability. To make an estimate of the degree of
spermiation failure, the number of spermatids retained in the
epithelium was compared with the number of spermatids available to
spermiate (Table 2). Animals that had
higher sperm counts had less retained spermatids and a lower percentage
of spermatids retained (Table 2). A significant negative correlation
was seen between sperm count and the percentage of spermatids retained
after 14 (r = -0.83, n = 5, P < 0.02) and
20 weeks (r = -0.94, n = 6, P < 0.001).
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Animals showed heterogeneous suppression of sperm counts, with some animals achieving consistent azoospermia, whereas others achieved variable degrees of spermatogenic suppression (17). As described elsewhere (17 ; and Narula et al., submitted), animals were assigned into two groups: those showing sustained azoospermia up to 20 weeks of T administration (azoo group, n = 4), and those that did not show consistent suppression, with consistently detectable sperm counts ranging between 1 and 40% of their control values (nonazoo group, n = 5).
Figure 4 shows germ cell development
(N/SC expressed as a percent of control) at 14 weeks of T
administration in the azoo and nonazoo groups. There was a general
trend for germ cell numbers to be lower in the azoo, compared with the
nonazoo group, from B spermatogonia onwards. When the data were
expressed as percent of control, significant decreases in the azoo
group were seen in pachytene spermatocytes, round spermatids, and steps
13–14 elongated spermatids. When the data were expressed as N/SC, germ
cell numbers were significantly lower in the azoo, compared with the
nonazoo, group in all germ cell groupings except for Ap spermatogonia
(not shown). There were no significant differences in the numbers of
germ cells between azoo and nonazoo groups before T administration (not
shown).
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shows the changes in the
spermatogonial subtypes in the azoo vs. nonazoo groups after
14 weeks of T administration. Animals in the azoo group had
significantly less Ad and B spermatogonia per Sertoli cell but similar
numbers of Ap. After 20 weeks, a similar pattern was observed, except
that there was no difference in the number of Ad spermatogonia between
the two groups (not shown).
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Although there was no difference in the levels of FSH between the azoo and nonazoo groups during the control phase, animals in the azoo group had significantly lower immuno-FSH and bioFSH across the 20-week T-administration phase, as detailed elsewhere (17 ; and Narula et al., submitted). After 14 weeks of T administration, bioFSH was 8% of control in the azoo group vs. 77% of control in the nonazoo group (P < 0.05). Similarly, after 20 weeks, bioFSH was 22% of control in the azoo group vs. 65% of control in the nonazoo group (P < 0.05). No differences were seen between the azoo vs. nonazoo groups, during the treatment period, in any other endocrine parameter, including serum T, bioLH, and intratesticular androgens (17 ; and Narula et al., submitted).
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Discussion |
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The arrest of spermatogenesis at the level of type A spermatogonia, in response to gonadotropin withdrawal, is supported by other studies in monkeys, after GnRH antagonist administration (9, 10, 11) and hypophysectomy (8), and in humans given weekly injections of T enanthate (15). In the current study, we showed that, whereas total A spermatogonial numbers were unchanged after 2 weeks of T administration, a decline in Ap spermatogonial number was apparent; and thus, Ap spermatogonia were the first germ cell type to show a significant decrease after gonadotropin withdrawal. In all animals, a major reduction in Ap spermatogonial number was seen at 14 and 20 weeks of T administration, and CR data showed that the progression of germ cells through the pachytene spermatocyte stage and spermiogenesis was also impaired, albeit to a lesser extent. We speculate that losses of pachytene spermatocytes and spermatids result from apoptosis, as has been previously described in gonadotropin-deprived rats (26, 27).
We recently reported that monkeys achieving consistent azoospermia had significantly lower levels of circulating FSH (both immuno- and bioactive FSH) (17 ; Narula et al., submitted), whereas no differences were seen in serum bioLH and T, or testicular androgen levels. These data suggest that residual FSH plays a key role in supporting spermatogenesis during T-induced gonadotropin suppression. The heterogenous sperm count response seen in these monkeys also allowed us to compare germ cell numbers, sperm counts, and FSH levels in animals achieving either consistent azoospermia or variable spermatogenic suppression. The 14-week time point during T administration was investigated to understand changes in the spermatogenic process that would promote subsequent changes in sperm output during the remaining 6 weeks of the T-administration phase. Analysis of the germ cell data at 14 weeks showed that, whereas Ap spermatogonia were similarly suppressed in both the azoo and nonazoo groups, all other germ cell types were lower in the azoospermic animals. Analysis of CR between germ cell types showed that the reduction in the transition of Ap to B spermatogonia was a major subsequent determinant of sperm output. Therefore, induction of consistent azoospermia is associated with lower circulating FSH levels (17 ; and Narula et al., submitted), a greater impairment of either B spermatogonial survival or in the transition of Ap to B spermatogonia, leading to fewer germ cells being available to proceed through spermatogenesis.
Type A spermatogonia in the primate and the human can be divided into at least two morphologically distinct subtypes: Ad and Ap (22). Type Ap spermatogonia divide in the later stages of the spermatogenic cycle, to produce type B spermatogonia, as well as to renew their own population (21, 22, 24, 25, 28, 29). Ad spermatogonia, however, rarely divide and are thus considered to be resting or reserve stem cells (21, 22, 24, 25, 28, 29). Ap spermatogonia have been suggested to be the true stem cells of the testis, because Ap (and not Ad) spermatogonia are seen in humans after radiation therapy (30), after long-term estrogen therapy, and in the postpubertal cryptorchid testes (31). Thus, Ap spermatogonia are probably the stem cells of the testis that renew their own population as well as produce cells committed to differentiation. Various studies have suggested that Ap spermatogonia can undergo a transition, probably without division, into Ad spermatogonia (24, 28, 29). Because Ad spermatogonia are not proliferative, the transition of Ap into Ad may be a means of inactivating a certain proportion of the spermatogonial population (28, 29). Moreover, studies in irradiated monkeys suggest that Ad spermatogonia may be able to undergo a transition back to Ap spermatogonia, thus allowing repopulation of the testis (28, 29), further suggesting that Ad spermatogonia are reserve stem cells.
T implants caused a decrease in Ap spermatogonia within 2 weeks, and this decrease in Ap spermatogonia eventually led to subsequent decreases in germ cells from type B spermatogonia onwards. Other investigators have shown reductions in Ap spermatogonia after long-term gonadotropin suppression (9) or hypophysectomy (8) in macaques. This reduction may involve the inhibition of the mitosis of Ap spermatogonia, because GnRH antagonist treatment of macaque monkeys decreased Ap spermatogonia in the stages in which mitosis occurs (11), and this led to a disappearance of proliferating cell nuclear antigen-labeled Ap spermatogonia from the testis (25). The relative role of androgens and FSH in this loss of Ap spermatogonia is unclear. FSH treatment stimulated Ap spermatogonial number in either GnRH antagonist-treated (9) or normal (12) macaques; however, their number can also be maintained by androgens (8), suggesting a role for each hormone. In the current study, Ap spermatogonia were maximally suppressed within 2 weeks, at a time when there were more substantial reductions in testicular T than in serum FSH. Our study paradigm does not allow us to identify the relative contributions of the suppression of T and FSH to the impairment of Ap spermatogonial number. Clearly, the fall in intratesticular T levels could have contributed to the reduction in Ap spermatogonia; however, we cannot exclude the possibility that the reduction in serum FSH bioactivity is the major regulator of this impairment.
Species differences between primates in the regulation of spermatogonial number may exist. In rhesus monkeys, B spermatogonia, rather than Ap spermatogonia, seem to be hormone-sensitive, because only B spermatogonial numbers increased after administration of FSH to hypophysectomized animals (13). Also, unilaterally orchidectomized rhesus monkeys, which had significant increases in circulating FSH levels, showed increases in B spermatogonia but not Ap populations (14). However, although a change in Ap numbers was not seen, it is possible that the mitotic activity of Ap spermatogonia may still be stimulated by FSH in these monkeys, leading to the increased numbers of B spermatogonia. In juvenile rhesus monkeys, FSH alone (32), or both FSH and T (33), stimulated Ap spermatogonial number, suggesting that, in young rhesus monkeys, these cells are hormonally responsive.
The lower circulating FSH levels in azoospermic animals were associated with a significant decrease in the conversion of Ap to B spermatogonia, leading to a lower number of B spermatogonia per Sertoli cell. Thus, in the azoospermic animals, B spermatogonia may have had a decreased mitotic activity and/or undergone apoptosis, because of lower levels of FSH. As the number of Ap spermatogonia were similarly reduced in both azoo and nonazoo groups, we speculate that Ap spermatogonia are maintained at a higher level of FSH than are type B spermatogonia. An alternative explanation is that the lower FSH levels in the azoospermic animals cause a further reduction in the mitotic activity of Ap spermatogonia during their differentiation into type B spermatogonia, leading to few B spermatogonia being produced. Therefore, all of the effects of FSH suppression may be at the level of Ap spermatogonia rather than B. A third explanation for the data is that Ap spermatogonia may be regulated by FSH and/or T (8, 9, 12), whereas B spermatogonia may be primarily regulated by FSH, as has been postulated (13).
The suggestion that Ap and B spermatogonia may be differentially regulated may explain the discrepancy between our current and previous studies (11). Though the current study shows that 2 weeks of T administration caused a decrease in Ap spermatogonia (yet no change in type B), previous studies in monkeys given a GnRH antagonist for 16 or 25 days demonstrated decreases in B spermatogonia in the absence of significant changes in Ap spermatogonia (11). Although that study showed Ap spermatogonial mitosis was impaired, as evidenced by a decrease in the proportion of Ap spermatogonia in stages VII-XII, wherein Ap spermatogonial mitosis occurs, the primary defect was in type B spermatogonia (11). FSH levels were not measured in the GnRH antagonist-treated animals (11), but it is possible that GnRH antagonist treatment caused a more rapid suppression of circulating FSH than T administration, leading to more marked and rapid effects on B spermatogonia.
The increase in Ad spermatogonia after 2 weeks of T administration, and a second increase after removal of the T implants, was unexpected. It is believed that Ad spermatogonia do not proliferate (22, 25), and thus they are unlikely to proliferate in response to T administration. We suggest that Ap spermatogonia cease to proliferate after gonadotrophin withdrawal, as has been demonstrated (25), and instead undergo a transition into resting Ad spermatogonia. Ap spermatogonia have been suggested to form Ad spermatogonia in normal monkeys via transitional type A (At) spermatogonia (24). In our study, At spermatogonia were classified as Ad spermatogonia. Studies in irradiated monkeys suggest that Ap are inactivated into Ad spermatogonia soon after irradiation (28, 29). Thus, it is possible that gonadotropins stimulate Ap proliferation, whereas gonadotropin suppression causes a transition into nonproliferative Ad forms. Studies in rhesus monkeys have shown decreases in Ad spermatogonia after either administration of FSH to juvenile monkeys (34) or a unilateral-orchidectomy-induced rise in FSH (14). Thus, Ad spermatogonia are increased during gonadotropin suppression, when there is a decrease in more differentiated spermatogonial subtypes (current study), yet are decreased during FSH stimulation, when more differentiated spermatogonia are present and proliferating (14, 34). The subsequent decline in Ad spermatogonia after 14 and 20 weeks of T administration in the current study may have been attributable to further losses of Ap spermatogonia or a decrease in the rate of transition into Ad forms. At 12 weeks of recovery, Ad spermatogonia were increased, and Ap spermatogonia decreased, yet the total population of A spermatogonia was normal. This data suggests that altered spermatogonial dynamics persist for at least 2 spermatogenic cycles.
The failure of spermiation was apparent after long-term T administration. There were few retained spermatids in biopsies taken after 2 weeks of treatment, and no significant reduction in sperm counts, suggesting that significant spermiation failure did not occur within 2 weeks. Although considerable spermiation failure was seen after 1 week of FSH and T suppression in rats, the experimental design ensured a rapid suppression of gonadotropin and T levels (16). In monkeys, spermatid retention has been noted after 16 and 25 days of GnRH antagonist treatment (11) and may be attributable to a more rapid suppression of gonadotropins with this treatment. Also, trials in humans, using T in combination with cyproterone acetate to promote the rapid suppression of gonadotropins, leads to a marked suppression of sperm counts within 4 weeks (3), suggesting early defects in spermiation. In the current study, spermiation failure was present, to a variable extent, after 14 and 20 weeks of T administration. Spermiation failure was particularly evident in animals with severe spermatogenic suppression, i.e. sperm counts less than 3 million per ejaculate. For example, one animal (no. 86185) had sperm counts which ranged from 0.1–1.5 million per ejaculate, germ cell numbers approximately 40% of control, and high numbers of retained spermatids, compared with animals with less severe suppression of sperm counts. Another animal (no. 97114) that was azoospermic between weeks 16–20 continued to show germ cell numbers at approximately 35% of control and had high numbers of retained spermatids. The demonstration of a negative correlation between sperm counts and retained spermatids suggests that spermiation failure contributes to a more severe suppression of sperm counts. The reason for spermiation failure in individual monkeys in the current study was not immediately obvious from their serum hormone data. Other studies involving long-term suppression of gonadotropins in monkeys have noted an accumulation of elongated spermatids in the testis, indicating spermiation failure (35). Our recent study in men receiving long-term T enanthate treatment also showed that some men who became azoospermic had elongated spermatids present in their biopsies, indicating that spermiation failure is a feature of long-term T administration in humans (15). Spermiation failure is seen after FSH and/or T suppression in rats (16) and is often noted in other situations, such as after the administration of reproductive toxicants (see 36 for review). The mechanism of spermiation failure is unknown but is presumably mediated by the Sertoli cell, because this cell contains the hormone receptors, and the elongated spermatids have ceased to be transcriptionally active. Immunocytochemical studies in rats have suggested that spermatids that fail to spermiate retain adhesion molecules on their surface, suggesting that a defect in the removal of adhesion molecules may be involved (37).
In summary, this study has described the effects of T-induced gonadotropin suppression on germ cell populations in adult macaques. Gonadotropin suppression caused a primary defect in Ap spermatogonia that leads to subsequent decreases in other germ cells. Cells were also lost during the pachytene spermatocyte and spermatid phases, probably via apoptosis. Spermiation failure was demonstrated in animals with marked suppression of sperm counts, yet maintenance of elongated spermatid production in the testis. Animals that had low circulating FSH with an induction of consistent azoospermia (17 ; and Narula et al., submitted) had a significant defect in the transition of Ap to B spermatogonia, suggesting an important role for FSH in the maintenance of B spermatogonia. The data suggests that, although defects in Ap spermatogonial development is a primary defect in monkey spermatogenesis, the basis for the variable suppression of sperm counts in T-treated monkeys is attributable to a more profound suppression of the transition between Ap and B spermatogonia, as well as defects in spermiation.
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Received October 4, 2000.
Revised December 4, 2000.
Accepted December 19, 2000.
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-reductase activity impairs
the testosterone-dependent restoration of spermiogenesis in adult rats. Endocrinology. 137:2703–2710.[Abstract]
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), type B
spermatogonia (B, •), preleptotene-zygotene spermatocytes (Pl-Z,
), pachytene spermatocytes (PS, 
). B, Data for spermatid
populations, steps 1–7 rSt (rSt 1–7, 
