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Prince Henry’s Institute of Medical Research (L.O., R.I.M., D.M.R.), Clayton, 3168 Victoria, Australia; and Department of Urology, Yokahama City University School of Medicine (K.S.), 3–9 Fukuura, Kanazawa-ku, Yokahama, Kanagawa 236, Japan
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, 3168 Victoria, Australia. E-mail: liza.odonnell{at}med.monash.edu.au
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Abstract |
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Introduction |
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It has been known for many years that the degeneration of step 19 spermatids during spermiation results from gonadotropin suppression due to hypophysectomy (3; see Ref. 4 for review). Many researchers have noted the appearance of degenerating step 19 spermatids in rats after hormone suppression (5, 6, 7, 8, 9, 10) and the administration of reproductive toxicants (11, 12, 13). A quantitative study by Russell and Clermont (14) demonstrated that rats hypophysectomized for 5.5 days had significant increases in the number of degenerating step 19 spermatids when expressed per Sertoli cell. This degeneration was prevented in part by supplementation with LH and was completely suppressed by supplementation with both FSH and LH. The researchers showed that germ cells in stage VII were susceptible to degeneration after hormone suppression, with step 19 spermatids being particularly vulnerable. These studies, however, did not provide an indication of the relative contribution of this degenerative process to the overall reduction of sperm production caused by these treatments.
When comparing the number of elongated spermatids in testicular biopsies to the sperm output in the ejaculates of men undergoing testosterone (T)-based contraception, we noted that all four men who became azoospermic had mature spermatids present (ranging from 1.4–20% of the control value) in the seminiferous epithelium (15). This finding suggested that spermiation failure may contribute to spermatogenic suppression induced by T-based contraception (15).
This study aimed to assess the extent to which spermiation failure contributes to the overall failure of spermatogenesis during hormone suppression. This necessitated a comparison between the number of normal spermatids before spermiation and the number of degenerating step 19 spermatids retained in the epithelium after stage VII. So that the two populations, which are present at different stages with different durations, could be compared, it was necessary to express the number of normal and retained spermatids produced per hour. We used the optical disector stereological approach, which is an assumption-free method (see Ref. 16 for review) for the quantitation of normal and retained spermatid nuclei in various stages on a per testis basis. This method has been used previously to quantitate elongated spermatid populations in the mouse, rat, and human (15, 17, 18).
Previous studies have investigated the degree to which spermatid degeneration after gonadotropin suppression by hypophysectomy can be prevented by supplementation with FSH and/or LH/T to understand which hormones regulate spermiation (14). We reexamined and extended these earlier studies by using in vivo models to specifically and acutely suppress either FSH or T. Acute suppression of FSH is achieved by passive immunization of adult rats with a FSH antibody, which has been shown to bind more than 90% of circulating FSH (19). Acute suppression of LH/T is achieved by pretreatment of rats for 1 week with 24-cm T (T24) implants to suppress serum LH yet maintain spermatogenesis, then implantation with low doses of T and estradiol (TE implants) to acutely suppress testicular T and thus spermatogenesis (20).
Thus, the following study aimed to use stereological methods to examine the effects of 1 week of FSH, T, or FSH plus T suppression on the proportion of spermatids that fail to spermiate. Further, we examined earlier spermatocyte and spermatid populations after T suppression between 0–6 weeks to understand how the failure of spermiation relates to losses in earlier germ cell types over the time course of spermatogenic failure. The results show that a high proportion of spermatids fail to spermiate after hormone suppression, and that this process may contribute to spermatogenic failure during T-based contraception.
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Materials and Methods |
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Steroid implants
T and estradiol (Sigma, St. Louis, MO)-filled
SILASTIC brand implants (Dow Corning Corp., Midland, MI)
were prepared using medical grade polydimethylsiloxane tubing
(Dow Corning Corp.; id, 1.98 mm; od, 3.18 mm), and medical
adhesive silicone type A as previously described (21). T implants were
either 3 or 24 cm (8 x 3 cm) in length, whereas estradiol
implants were 0.4 cm in length.
Rat FSH polyclonal antibody
The Ig fraction of a sheep polyclonal antiserum raised against
NIDDK I-8 rat FSH has been previously characterized (FSHAb) (19). The
Ig fraction of serum from nonimmunized sheep (ConAb) was used as the
control. The administration of the FSH antibody for 1 week to adult
rats has been previously shown to immunoabsorb at least 90% of
circulating FSH (19). The FSHAb and the ConAb were administered sc at a
dose of 2 mg/kg·day in sterile 0.154 M NaCl.
Experimental design
Exp 1: 1 week of T and/or FSH withdrawal. Twenty-four rats
were used for Exp 1. Twelve animals received 24-cm T implants sc for 1
week to maintain spermatogenesis but suppress circulating LH, as
previously described (20). This procedure ensured that serum LH was
fully suppressed before low dose T and estradiol treatment. The T24
implants were then removed and replaced with 3-cm T and 0.4-cm
estradiol implants (TE) for 1 week to suppress testicular T and
spermatogenesis (20). During the 1 week of TE treatment, half of the
animals received daily injections of ConAb (T alone withdrawal), and
the other half of the animals received daily injections of FSHAb (FSH+T
withdrawal). Six animals received no implants, but were given the same
dose of FSHAb daily for 1 week (FSH alone withdrawal). This passive
immunization was limited to 1 week due to antibodies being produced to
the exogenously administered IgG (19). Six animals received no
implants and daily injections of ConAb, and served as the control
group.
Exp 2: time course of T withdrawal. Animals received 24-cm T implants for 1 week as described above, and then implants were removed and replaced with TE implants. Animals were killed 0, 1, 2, 3, 4, 5, and 6 weeks after TE implantation (n = 6/group). This procedure was thus an extension of the TE treatment as explained above. Control animals (n = 6) did not receive any implants (either T24 or TE). The LH and testicular T levels in animals receiving T24 plus TE implants for 0–6 weeks have been previously described (20).
Tissue collection
Animals received 100 IU heparin sc 0.5–2 h before perfusion.
Immediately after ether anesthesia, blood was collected by cardiac
puncture, and plasma was stored at -20 C for hormone assays. The right
testis was then excised and weighed. For perfusion fixation, the dorsal
aorta was cannulated, and the vasculature was flushed with 0.154
M NaCl before fixation with Bouin’s fixative, as
previously described (21). The testis perfused in situ was
then removed, subjected to systematic uniform random sampling (16), and
embedded in methacrylate resin, and 25-µm periodic
acid-Schiff-stained sections were prepared as previously described
(22).
Cell number estimates
The optical disector method (15, 19, 22; see Ref. 16 for review)
was used to determine the numbers of germ cells and retained spermatids
per testis. Twenty-five-micron-thick methacrylate sections were
optically sectioned by means of a high numerical aperture lens (x100
objective lens, N.A. 1.4). The Olympus Corp. BX-50
microscope (Tokyo, Japan) was equipped with a microcater (D83301,
Heideinhain, 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). Germ cells were
identified using previously described morphological criteria (23). In
each field, a counting frame (area 625 µm2) was
used to count germ cells, whereas a larger frame (area 3125
µm2) was used to count retained spermatids.
More than 600 fields/animal were evaluated. Thus, the total area
assessed for each testis was more than 1.875 mm2.
The final screen magnification was x2708. As previously determined, no
correction for tissue shrinkage was necessary (24).
Morphologically normal germ cells were counted in the following
categories: pachytene spermatocytes in stages I–III, IV–VI, VII,
VIII, and IX–XIII (diplotene spermatocytes were included in this
grouping); round spermatids in stages I–VI (steps 1–6 spermatids),
VII (step 7 spermatids), and VIII (step 8 spermatids); and elongated
spermatids in stages IV–VI (step 17–18 elongated spermatids). Data
from the various stage groupings were pooled for purposes of
presentation where applicable. Germ cell numbers were calculated on a
per testis basis (19, 22) and then divided by the stage duration in
hours, using published time divisors (25) to express cell numbers in
millions per hour. The conversion of step 7 to 8 round spermatids was
determined by dividing the number of round spermatids in stage VIII in
millions per hour by the number of round spermatids in stage VII in
millions per hour (20, 21, 26). For each animal, more than 80 nuclei of
each germ cell type in each stage grouping were counted. However, in
some treatments (T withdrawal for 4–6 weeks) step 8 round spermatids
and step 17–18 spermatids were scarce, and fewer cells (
20–40)
were counted.
Quantitation of retained spermatids
Retained spermatid nuclei were counted using the stereological
procedures described above. A retained spermatid was defined as a
curved elongated nucleus 5 µm or more in length situated in the basal
and intermediate portions of the epithelium, and these were counted in
each of stages VIII–XIII. In hormone-treated animals, at least 50
(usually >100) retained spermatids were counted in each stage.
However, in control animals few spermatids were seen, and only 10
cells/stage could be evaluated. The number of retained spermatids per
testis was calculated as described above. The number of retained
spermatids was also expressed in millions per hour by dividing the
number per testis by the relevant stage duration in hours (25).
The percentage of spermatids failing to spermiate was calculated by dividing the number of retained spermatids in stage IX in millions per hour (see Results) by the number of step 17–18 spermatids in millions per hour, multiplied by 100. The percentage of spermatids failing to spermiate was not calculated when the number of step 17–18 spermatids was less than 0.1 x 106/h because of difficulties associated with substaging stages IX–XIII when spermatogenesis is markedly suppressed. Therefore, no such calculation was performed in the TE 6 week group, as step 17–18 spermatids were close to undetectable levels. Also, two animals in the TE 4 week group and three animals in the TE 5 week group had few step 17–18 spermatids and were therefore excluded from the analysis.
FSH RIA
Serum FSH was determined by double antibody RIA as previously
described (21). The FSH RIA used iodinated rat (r) FSH (NIDDK,
Bethesda, MD; rFSH I7) as tracer, rFSH antiserum (NIDDK anti-rFSH S11),
and rFSH reference preparation 2 as standard. All samples were measured
in one assay, with a sensitivity of 1.3 ng/ml. FSH levels were not
determined in Exp 1 due to the interference from the administered FSH
antibody.
Statistics
In Exp 1, differences among groups were assessed by ANOVA and
Peritz’s multiple range test at the level of P <
0.05. In Exp 2, differences compared with the control group were
assessed by ANOVA and Student’s t test at the level of
P < 0.05 unless otherwise stated. All data are
expressed as the mean ± SEM (n =
6/group).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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. In all groups, the number of retained
spermatids peaked in stage IX and then gradually declined. Given that
in both control and treatment groups the maximum number of retained
spermatids was detected in stage IX, the population of retained
spermatids in stage IX was considered to represent the entire
population of retained spermatids before their presumed digestion by
Sertoli cells. Therefore, the percentage of spermatids failing to
spermiate was calculated as the number of retained spermatids in stage
IX (millions per hour) divided by the number of step 17–18 spermatids
(millions per hour), multiplied by 100.
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Germ cell populations. After 1 week of T, FSH, or FSH plus T
withdrawal (see Fig. 2), there were no
changes in the populations of stage I–XIII pachytene spermatocytes
(Fig. 2A) or step 17–18 spermatids (Fig. 2C). There was a small, but
significant, decrease in the number of step 1–8 round spermatids after
1 week of FSH plus T withdrawal (Fig. 2B).
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Numbers of retained spermatids. The total numbers of retained
spermatids in stages VIII–XIII in control animals and in those animals
in which FSH, T, or FSH plus T were suppressed for 1 week are shown in
Fig. 3A. In control animals, 0.74 ±
0.11 million retained spermatids/testis were observed in stages
VIII–XIII. In response to FSH alone or T alone suppression for 1 week,
the numbers of retained spermatids per testis increased to 5.5- and
6.4-fold that in control animals (P < 0.05),
respectively. There was no significant difference in the number of
retained spermatids between FSH alone and T alone withdrawal. When FSH
and T were withdrawn in combination, there was a marked increase in the
number of retained spermatids, to 35.7-fold that in control
animals.
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. The percentage failing to spermiate was 2.3 ± 0.7% in
control animals. After 1 week of FSH or T withdrawal, 13.7 ±
4.3% and 10.9 ± 1.9% of elongated spermatids failed to
spermiate, respectively. However, suppression of FSH and T in
combination for 1 week increased the percentage failing to spermiate to
49.7 ± 5.0%.
Exp 2: time course of T withdrawal
Testis weight. TE treatment caused a progressive decline in
testis weight, which became significant by 2 weeks (1.48 ±
0.05 g; control, 1.77 ± 0.01 g; P <
0.05) and reached 0.55 ± 0.04 g (P < 0.05)
by 6 weeks of TE treatment.
Serum FSH levels. Treatment with T24 followed by TE caused a transient significant decrease (P < 0.05) in serum FSH in the TE 3 week (5.6 ± 1.8 ng/ml) and TE 4 week (6.0 ± 1.6 ng/ml) groups compared with the control value (9.2 ± 1.4 ng/ml). Other groups were not different from the control (data not shown).
Germ cell populations. The numbers of germ cells during the
time course of T suppression are shown in Fig. 4. Pachytene spermatocyte numbers (Fig. 4A) were divided into stages I–VII and VIII–XIII to investigate the
loss of early spermatocytes with time as well as to represent midcycle
losses of pachytene spermatocytes. The decline in stage I–VII
pachytene spermatocytes occurred gradually, becoming significantly
different from the control value after 3 weeks and reaching 54.4% of
the control value by 6 weeks of treatment (P < 0.05).
Stage VIII–XIII pachytene spermatocytes also declined over the time
course, becoming significantly different by 2 weeks and reaching 31.4%
of the control value by 6 weeks. Furthermore, a significant decrease
(P < 0.05) in pachytene spermatocytes in stages
VIII–XIII compared with pachytene spermatocytes in stages I–VII was
noted by 2 weeks of TE treatment, and this decrease was seen throughout
the rest of the treatment period.
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. The conversion
ratio was significantly decreased by 3 weeks of TE treatment and
reached 23.7% of the control value by 6 weeks.
The number of elongated spermatids before spermiation is shown in Fig. 4C. The number of step 17–18 spermatids was significantly suppressed
compared with the control value by 2 weeks. There was a marked loss of
elongated spermatids between 3 and 4 weeks. By 6 weeks of T
suppression, step 17–18 spermatids were reduced to 0.4% of the
control value.
Numbers of retained spermatids. The total number of retained
spermatids per testis during the time course of T suppression is shown
in Fig. 5A. There was a small, but
significant, increase in the number of retained spermatids after
pretreatment with T24 implants for 1 week (TE 0 week). Spermatid
retention was maximal after 2 weeks of TE treatment, and thereafter the
number of retained spermatids declined, until none was seen by 6
weeks.
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. By 2 weeks of T suppression, 45.2
± 11.4% of spermatids failed to spermiate, and this percentage
increased with increasing time of T withdrawal. It should be noted that
the percentage of spermatids failing to spermiate was not calculated in
animals that produced few step 17–18 spermatids (see Materials
and Methods). After 4 and 5 weeks of T withdrawal, when step
17–18 spermatids were being produced, the majority (>90%) of these
spermatids failed to spermiate. |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The majority of retained spermatids were found in stage IX, and the decline in their number in subsequent stages is attributed to their phagocytosis and digestion by Sertoli cells. Retained spermatid nuclei seen in stages VIII and IX appeared similar to normal elongated spermatid nuclei in stages VII and VIII. However, as phagocytosis by the Sertoli cell progressed, the retained nuclei appeared smaller and finer in stages X–XIII. Fragments of nuclei remained in the epithelium in stages XIV and I, but these were not counted due to uncertainty in their identification. Using the optical disector method to quantitate the production rates of step 17 and 18 spermatids, we were able to relate this number to that of retained spermatids. This calculation, which used the number of retained spermatids in stage IX in millions per hour as the numerator and the number of steps 17–18 spermatids in millions per hour as the denominator, makes the assumption that all retained spermatids will be detected in stage IX. It is possible that a proportion of spermatids could have been very rapidly digested before this, and therefore would not have been counted in stage IX. In this case the reported percentage of spermatids failing to spermiate would actually have been underestimated.
The suppression of either FSH or T for 1 week led to 11–14% of spermatids failing to spermiate. However, the combined suppression of FSH and T for 1 week led to a striking degree of spermiation failure, with half of the mature spermatids being retained. This marked increase in spermiation failure took place at a time when there were few changes in the number of earlier germ cells. We counted morphologically normal germ cells after 1 week of FSH and T withdrawal and saw minimal changes in earlier germ cell populations, such as spermatocytes and round spermatids in the hormone-sensitive stages VII and VIII. However, we did not enumerate degenerating germ cells in stages VII and VIII, and it is likely that there would have been significant increases in the number of degenerating spermatocytes and spermatids at these stages, as shown by others (8, 9, 14). Thus, other germ cells in stages VII and VIII will degenerate after 1 week of hormone withdrawal (14), although this loss produces no significant change in the viable germ cell population as enumerated by the optical disector.
The synergistic effect of combined FSH and T withdrawal on spermiation failure is supported by numerous studies showing synergy between FSH and T. The androgen requirements of spermatogenesis are much lower in the presence of circulating FSH (27, 28, 29, 30), suggesting that FSH may potentiate the action of T. Alternatively, the stimulatory effect of FSH on spermatogenesis may be potentiated by the presence of T, as has been suggested by studies in which the effect of FSH on the maintenance of germ cell populations in GnRH-antagonist treated rats was diminished when the antiandrogen, flutamide, was added (31). Furthermore, FSH and T have been suggested to have similar pathways of action with regard to the maintenance of cell viability (32). Presumably, suppression of both FSH and T removes the compensatory mechanisms that are in place when either one is depleted in isolation, and suppresses both signals which stimulate identical pathways to promote spermiation.
Exp 2 investigated the time course of T withdrawal on spermiation. One week of T24 treatment (i.e. TE zero time point) caused no change in the number of pachytene spermatocytes or round and elongated spermatids before spermiation, yet a significant increase in the number of retained spermatids was observed further emphasizing the sensitivity of spermiation to T suppression. Previously, T24 implants have been shown to maintain the number of homogenization-resistant spermatids in the testis at near-normal levels (28, 33); however, this method of quantitating elongated spermatids may or may not include retained spermatids. After 1 week of further T withdrawal by TE implants, failure of spermiation was again the only significant change in spermatogenesis. By 2 weeks of TE, there was a peak in the number of retained spermatids to approximately 25 million/testis, and approximately 50% of spermatids failed to be released. These figures are equivalent to the failure of spermiation after 1 week of FSH and T withdrawal in combination, suggesting that the depletion of FSH accelerates the time course of spermiation failure. Also, by 2 weeks of T withdrawal, there were decreases in the numbers of step 17–18 spermatids and pachytene spermatocytes in stages VIII–XIII, presumably due to the earlier degeneration of germ cells as they pass through hormone-sensitive stage VII (5, 14). By 3 weeks of TE there were further reductions in pachytene spermatocyte and step 17–18 spermatids, and a significant decrease in the conversion of round spermatids between steps 7 and 8, related to the induction of round spermatid sloughing (20). At this time, more than 70% of the spermatids available for spermiation were retained within the epithelium. By 4 and 5 weeks of TE there was a progressive loss of pachytene spermatocytes, which also reflects losses in earlier germ cell populations (33), further reductions in the ratio of step 7 to 8 round spermatids, and a marked decline in step 17–18 spermatids. Although the absolute number per testis of retained spermatids declined, fewer spermatids were progressing through the elongation process, and thus the percentage of spermatids failing to be released increased to more than 90%. By 6 weeks of TE, pachytene spermatocyte populations reached maximal suppression (33) as did round spermatid sloughing (20, 26), and due to round spermatid sloughing and germ cell degeneration, the number of step 17–18 spermatids was close to zero. At this time, very few retained spermatids were seen, highlighting the fact that spermiation failure is an early event during spermatogenic suppression.
In our previous study the basis for the delay until 3 weeks of TE treatment before round spermatids detached from the Sertoli cell was unclear (20). Given that the current study shows that round spermatid sloughing is preceded by a significant failure in spermiation, it is tempting to speculate that the two events are related. Recycling of ectoplasmic specializations from step 19 spermatids to newly formed step 8 spermatids has been suggested (1). However, retained spermatids do not appear to have ectoplasmic specializations attached to them, as evidenced by electron microscopy (14) and immunocytochemical localization of actin (2) and vinculin (2) (our unpublished observations); thus, it is possible that ectoplasmic specializations continue to be recycled normally. In agreement with this, we recently showed that ectoplasmic specializations were present in the seminiferous epithelium of the long-term TE-treated rat (34), further suggesting that the formation and recycling of these structures remain relatively normal. Another way in which spermiation may promote the progression of round spermatids through the elongation phase of spermiogenesis is via the normal phagocytosis of residual bodies. Indeed, residual bodies are thought to communicate with the Sertoli cell to trigger the release of various factors (35, 36). It is possible that spermiation may provide positive signals to the Sertoli cell to continue with spermiogenesis, and when this signal gradually declines to critical levels somewhere between 2–3 weeks of T withdrawal, round spermatid sloughing begins. Such hypotheses require further investigation.
Despite the fact that spermiation failure is noted under various conditions, surprisingly little is known about the molecular mechanisms involved in normal spermiation or the causes of spermiation failure (see Ref. 4 for review). Electron microscopic analysis of mature spermatids after hypophysectomy in rats suggested that there were abnormalities in the spermatid flagella, specifically that these flagellae were often enclosed in a thin sleeve of Sertoli cell cytoplasm, and that these abnormalities may have contributed to spermatid retention (14). Recent studies suggested that retained spermatids that were induced by boric acid treatment were positive for N-cadherin and desmoglein immunostaining (2). Although the molecular mechanisms of failure of spermiation after hormone withdrawal are unknown, the response is presumably mediated via the Sertoli cell, which contains the receptors for both FSH and T. Given that numerous toxicants can promote spermatid retention, failure of spermiation may be a nonspecific reaction to injury to the Sertoli cell.
In the human, it takes 64 days for spermatogonia to develop into mature spermatids (37). A feature of current male contraceptives is a slow onset of azoospermia and a slow recovery time. As spermiation is the final step of spermatogenesis, contraceptives that target spermiation would be advantageous in that they would allow a more rapid onset and recovery. Recent studies by Zhengwei and colleagues (15) used stereological techniques to evaluate germ cell populations in five men undergoing long-term T-based contraception. The data showed that although the most consistent defect in spermatogenesis in all five men appeared to be in spermatogonial maturation, there was also convincing evidence for a defect in sperm release. Data reported by Meriggiola and colleagues for men receiving T enanthate in combination with cyproterone acetate showed profound decreases in sperm counts within 4 weeks (38), suggesting significant defects in spermatogenesis at sites later than spermatogonial maturation. More studies on the hormonal sensitivity of spermiation in monkeys and humans as well as mechanistic studies on spermiation may eventually lead to contraceptive strategies that primarily target sperm release.
In summary, we have used stereological techniques to quantify the failure of spermiation that is induced by hormone withdrawal in adult rats. We show that although the acute withdrawal of either FSH or T produces similar degrees of spermatid retention, the combination of FSH and T withdrawal has synergistic effects, in that half of the mature spermatids in the testis fail to be released by 1 week. We show that spermiation failure is an early feature of spermatogenic suppression and has a significant contribution to the failure of spermatogenesis induced by hormone withdrawal. Our findings suggest that spermiation is a possible target for contraception.
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Acknowledgments |
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Footnotes |
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Received March 20, 2000.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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-reductase activity impairs
the testosterone-dependent restoration of spermiogenesis in adult rats.
Endocrinology 137:2703–2710[Abstract]
(IL-1) release, which triggers IL-6
production by an autocrine mechanism, through the lipoxygenase pathway.
Endocrinology 136:3070–3078[Abstract]
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 P. P Y Lie, W. Xia, C. Q F Wang, D. D Mruk, H. H N Yan, C.-h. Wong, W. M Lee, and C Y. Cheng Dynamin II interacts with the cadherin- and occludin-based protein complexes at the blood-testis barrier in adult rat testes J. Endocrinol., December 1, 2006; 191(3): 571 - 586. [Abstract] [Full Text] [PDF]
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K. L. Matthiesson, R. I. McLachlan, L. O'Donnell, M. Frydenberg, D. M. Robertson, P. G. Stanton, and S. J. Meachem The Relative Roles of Follicle-Stimulating Hormone and Luteinizing Hormone in Maintaining Spermatogonial Maturation and Spermiation in Normal Men J. Clin. Endocrinol. Metab., October 1, 2006; 91(10): 3962 - 3969. [Abstract] [Full Text] [PDF]
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A. Beardsley, D. M Robertson, and L. O'Donnell A complex containing {alpha}6{beta}1-integrin and phosphorylated focal adhesion kinase between Sertoli cells and elongated spermatids during spermatid release from the seminiferous epithelium. J. Endocrinol., September 1, 2006; 190(3): 759 - 770. [Abstract] [Full Text] [PDF]
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K. L. Matthiesson and R. I. McLachlan Male hormonal contraception: concept proven, product in sight? Hum. Reprod. Update, July 1, 2006; 12(4): 463 - 482. [Abstract] [Full Text] [PDF]
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P Sluka, L O'Donnell, J R Bartles, and P G Stanton FSH regulates the formation of adherens junctions and ectoplasmic specialisations between rat Sertoli cells in vitro and in vivo. J. Endocrinol., May 1, 2006; 189(2): 381 - 395. [Abstract] [Full Text] [PDF]
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S. J Meachem, D. M Robertson, N. G Wreford, R. I McLachlan, and P. G Stanton Oestrogen does not affect the restoration of spermatogenesis in the gonadotrophin-releasing hormone-immunised adult rat J. Endocrinol., June 1, 2005; 185(3): 529 - 538. [Abstract] [Full Text] [PDF]
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M Vigier, M Weiss, M H Perrard, M Godet, and P Durand The effects of FSH and of testosterone on the completion of meiosis and the very early steps of spermiogenesis of the rat: an in vitro study J. Mol. Endocrinol., December 1, 2004; 33(3): 729 - 742. [Abstract] [Full Text] [PDF]
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 M. Nistal, P. Gonzalez-Peramato, and R. Paniagua Diagnostic Value of Differential Quantification of Spermatids in Obstructive Azoospermia J Androl, September 1, 2003; 24(5): 721 - 726. [Abstract] [Full Text] [PDF]
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 J. Killian, K. Pratis, R. J. Clifton, P. G. Stanton, D. M. Robertson, and L. O'Donnell 5{alpha}-Reductase Isoenzymes 1 and 2 in the Rat Testis During Postnatal Development Biol Reprod, May 1, 2003; 68(5): 1711 - 1718. [Abstract] [Full Text] [PDF]
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A. Beardsley and L. O'Donnell Characterization of Normal Spermiation and Spermiation Failure Induced by Hormone Suppression in Adult Rats Biol Reprod, April 1, 2003; 68(4): 1299 - 1307. [Abstract] [Full Text] [PDF]
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W. W. Wright, L. Smith, C. Kerr, and M. Charron Mice That Express Enzymatically Inactive Cathepsin L Exhibit Abnormal Spermatogenesis Biol Reprod, February 1, 2003; 68(2): 680 - 687. [Abstract] [Full Text] [PDF]
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 D. Kinniburgh, H. Zhu, L. Cheng, A.T. Kicman, D.T. Baird, and R.A. Anderson Oral desogestrel with testosterone pellets induces consistent suppression of spermatogenesis to azoospermia in both Caucasian and Chinese men Hum. Reprod., June 1, 2002; 17(6): 1490 - 1501. [Abstract] [Full Text] [PDF]
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R. W. Bailey, B. Aronow, J. A.K. Harmony, and M. D. Griswold Heat Shock-Initiated Apoptosis Is Accelerated and Removal of Damaged Cells Is Delayed in the Testis of Clusterin/ApoJ Knock-Out Mice Biol Reprod, April 1, 2002; 66(4): 1042 - 1053. [Abstract] [Full Text] [PDF]
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R. I. McLachlan, L. O'Donnell, P. G. Stanton, G. Balourdos, M. Frydenberg, D. M. de Kretser, and D. M. Robertson Effects of Testosterone Plus Medroxyprogesterone Acetate on Semen Quality, Reproductive Hormones, and Germ Cell Populations in Normal Young Men J. Clin. Endocrinol. Metab., February 1, 2002; 87(2): 546 - 556. [Abstract] [Full Text] [PDF]
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 R.I. McLachlan, L. O'Donnell, S.J. Meachem, P.G. Stanton, D.M. de Kretser, K. Pratis, and D.M. Robertson Identification of Specific Sites of Hormonal Regulation in Spermatogenesis in Rats, Monkeys, and Man Recent Prog. Horm. Res., January 1, 2002; 57(1): 149 - 179. [Abstract] [Full Text] [PDF]
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L. O’Donnell, A. Narula, G. Balourdos, Y.-Q. Gu, N. G. Wreford, D. M. Robertson, W. J. Bremner, and R. I. McLachlan Impairment of Spermatogonial Development and Spermiation after Testosterone-Induced Gonadotropin Suppression in Adult Monkeys (Macaca fascicularis) J. Clin. Endocrinol. Metab., April 1, 2001; 86(4): 1814 - 1822. [Abstract] [Full Text]
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), T alone (
), FSH plus T (
), or no hormone (control; •).
Each point represents the mean ± SEM
(n = 6).
, data for treated animals. C, The number of step 17–18
elongated spermatids in millions per hour. All data are expressed as
the mean ± SEM (n = 6). *, P
< 0.05 compared with control; 
, P < 0.05
between stage I–VII and stage VIII–XIII pachytene spermatocytes.