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From the * Monash Institute of Medical Research,
Monash University, and Prince Henry's
Institute of Medical Research, Monash Medical Centre, Melbourne,
Australia.
| Correspondence to: A/Prof Mark Hedger, Monash Institute of Medical Research, Monash University, 27-31 Wright Street, Clayton, Victoria 3168, Australia (e-mail: mark.hedger{at}med.monash.edu.au). |
| Received for publication May 23, 2006; accepted for publication September 11, 2006. |
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Abstract |
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Key words: Apoptosis, meiosis, Sertoli cell, inflammation, fertility
Spermatogenesis takes place in the seminiferous tubules of the testis,
under the control of the gonadotropins, luteinizing hormone (LH) and
follicle-stimulating hormone (FSH), testicular androgens (testosterone), and
local regulatory networks. Spermatogenesis itself is a complex and highly
ordered process that involves progression of the seminiferous epithelium
through a series of germ cell associations called the cycle of the
seminiferous epithelium (Clermont,
1972). It is particularly significant that certain inflammatory
mediators appear to be involved in the coordinated regulation of this cycle.
The Sertoli cells, which support the developing germ cells in the epithelium,
produce the cytokines interleukin-1
(IL-1), IL-6, and the
transforming growth factor-ß family member activin A
(Stéphan et al, 1997;
Okuma et al, 2005). A complex
interaction between these cytokines within the seminiferous epithelium has
been implicated in cyclical regulation of spermatogonial proliferation and the
progression of meiosis (Mather et al,
1990; Parvinen et al,
1991; Hakovirta et al,
1995; Okuma et al,
2005). During spermatogenesis, programmed cell death (apoptosis)
plays an important role in modulating spermatogenic production capacity and
possibly in the elimination of defective germ cells. Tumor necrosis
factor- and Fas ligand are 2 locally produced cytokines that have been
implicated in controlling germ cell apoptosis in both normal and pathological
conditions (Boekelheide et al,
1998; Pentikäinen et al,
2001). Clearly, up-regulation of inflammatory processes that
involve these various cytokines will impinge upon the testis and have profound
and specific effects on various aspects of spermatogenesis. Moreover,
inflammatory cytokines exert mostly negative effects on Leydig cell function,
interfering with the production of the testosterone that is essential for
maintenance of spermatogenesis and male reproductive function in general
(Hales et al, 1999).
Lipopolysaccharide (LPS), a major component in the cell wall of gram-negative bacteria, has been widely used in studies on the effects of inflammation in many systems. At high doses, LPS causes septic shock, but at lower doses it induces a well-defined inflammatory state (Rietschel et al, 1991; Galanos and Freudenberg, 1993). In previous studies, disorganization of the seminiferous epithelium has been observed in adult rats treated with a sublethal dose of LPS (Tulassay et al, 1970; O'Bryan et al, 2000). This disorganization involves an increase in apoptosis and disruption of the essential cell-cell contacts between Sertoli cells and spermatocytes or spermatids, resulting in these germ cells being prematurely released into the tubule lumen (O'Bryan et al, 2000). However, the precise lesion of spermatogenesis and its timing has not been identified, nor have the longer-term consequences for fertility been assessed. Such information should provide vital information regarding the mechanisms involved in damage to fertility during inflammation. In order to investigate this further, we have undertaken a detailed stereological analysis of the spermatogenic process during and subsequent to an episode of acute LPS-induced inflammation. In this study, the contribution of changes in hormones was also examined through the measurement of serum gonadotropins, serum and intratesticular testosterone, and the Sertoli cell hormone inhibin B.
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Methods and Materials |
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Experimental Design
In the short-term study, rats were randomly assigned to the control group
or 1 of 3 treatment groups, each containing 7 animals. Treated rats received a
single intraperitoneal (IP) injection of LPS (from Escherichia coli,
serotype 0127:B8) (Sigma-Aldrich, St Louis, Mo) at a dose of 5 mg/kg body
weight and were killed at 1, 3, and 7 days after injection. Control groups
received an injection of an equivalent volume of endotoxin-free saline, and
were killed in groups of 2 or 3 on days 1, 3, and 7 after injection. For the
long-term study, rats were randomly assigned to a single LPS-treated (5 mg/kg
body weight) or saline-injected control group (7 animals/group) and were
killed 28 days later. Rats were observed throughout the experimental period
and general health conditions were noted and recorded.
Tissue Collection and Preparation
Rats were anesthetized with isoflurane USP inhalation anesthetic and the
right testes were excised, snap-frozen in liquid nitrogen, and stored at
–80°C for preparation of testicular homogenates, prior to whole-body
perfusion fixation. The chest was cut open to gain access to the heart, and 5
to 10 mL blood was collected by cardiac puncture. Blood was then centrifuged
(15 min, 3000 x g) at room temperature and the serum stored at
–20°C for subsequent hormone analysis. The descending aorta was
cannulated, and the vasculature was flushed with 0.9% saline prior to
perfusion fixation with Bouin fluid for 5 minutes. Following perfusion, the
left testis was excised and placed in Bouin fixative for no more than 5 hours.
The left testis was then weighed and sampled for stereological analysis in a
systematic uniform random sampling manner
(Wreford, 1995). The testis
was sliced into a series of 2-mm parallel discs orthogonal to the long axis
(8–9 discs in each testis). Three discs per animal were selected, and
half of each was processed and embedded into hydroxethyl methacrylate resin
(Technovit 7100 methacrylate resin kit; Kulzer and Company GmbH,
Friedrichsdorf, Germany) according to the manufacturer's instructions while
the other half was used for routine embedding into paraffin. Thick (25-µm)
resin sections were serially cut (2050 Supercut microtome; Reichert-Jung,
Nossloch, Germany) and stained with periodic acid-Schiff (PAS) reaction
reagents for histology as previously described
(Meachem et al, 1997). Thin
(5-µm) paraffin sections were transferred to Superfrost Plus slides for
analysis of testicular apoptosis and histology.
Testicular Homogenates
Testis tissue (0.5 g) was hand-homogenized in 3 volumes of ice-cold
phosphate-buffered saline (pH 7.4) and ultracentrifuged (105 000 x
g, 70 min, 4°C). The supernatants containing the testicular
cytosol fraction were collected and stored at –20°C prior to
assay.
Stereological Analyses
The optical dissector method (Wreford,
1995) was used to determine the total number of Sertoli and germ
cells per testis. All estimates were performed using a 100x objective on
an Olympus BX-50 microscope (Tokyo, Japan). The images were captured by a JVC
TK-C1381EG (JVC, Yokohama, Japan) color video camera coupled to a Pentium PC
computer. A software program, DH CASTGRID V1.10 (Olympus, Munich, Germany) was
used to superimpose a set of unbiased counting frames on the video image.
Fields were selected using a systematic uniform random sampling scheme as
previously described (McLachlan et al,
1994; Wreford,
1995), generated using a computer-driven motorized stage
(Multicontrol 2000; ITK, Lahnau, Germany). A microcator (Heidenhain D83301;
Heidenhain, Traunreut, Germany) was attached to the microscope stage and
monitored depth measurement.
The Sertoli and germ cells were identified based on their location within the cord, their size, their acrosomal level, and the shape of the cell nucleus (Russell et al, 1990). The frame size for counting the Sertoli cell, type A spermatogonia (associated with stages I–XIV), type B spermatogonia (stages V–VI), preleptotene (stages VII–VIII), and leptotene and zygotene spermatocytes (stages IX–XIII) was 2820 µm2. On the other hand, 4 frames with a total of 940 µm2 frame size were used to count pachytene spermatocytes (associated with stages I–VIII and IX–XIV) and 1 out of the 4 frames for round spermatids (associated with steps 1–8), elongating spermatids (associated with steps 9–14), and elongated spermatids (associated with steps 15–19). The difference in the number of fields and frame sizes depended on the frequency of the cells observed. Slides were masked prior to each type of quantification (cell number and apoptosis as outlined below) to facilitate unbiased counting.
Histological Analyses
The paraffin sections (5 µm) were stained with PAS for general
histology, as modified from Meachem et al
(1997). Sections were
deparaffinized and rehydrated in a series of graded concentrations of ethanol.
The sections were immersed in a solution of 0.5% periodic acid for 10 minutes.
The slides were washed in running tap water and immersed in Schiff reagent
(Amber Scientific, Belmont, Australia) for 10 minutes, counterstained with
Harris hematoxylin (Sigma Aldrich, St Louis, Mo), dehydrated in graded
ethanol, and mounted under glass coverslips using DPX (BDH Laboratory
Supplies, Poole, United Kingdom).
In Situ Detection of DNA Fragmentation
Tissue sections (5 µm) were deparaffinized and rehydrated prior to the
detection of DNA fragmentation. Apoptotic cells were detected using the
terminal deoxynucleotidyl transferase (TdT)-mediated dUTP-biotin nick-end
labeling (TUNEL) method, modified from Meachem et al
(1999). On negative control
sections, the TdT enzyme was omitted.
Three sections per rat were examined for the presence of TUNEL-positive germ cells in controls and LPS-treated rats. Each tubule cross-section was classified into 3 stage groupings (stages XIV–IV, V–VIII and IX–XIII). TUNEL-positive cells were identified by dark brown staining. In order to estimate number of TUNEL-positive cells per tubule cross-section, a total of between 90 and 100 tubules for each stage grouping were studied per rat. All slides were masked prior to the study.
Hormone Assays
Concentrations of serum FSH levels were determined by using an
immunofluorometric assay (IFMA) as previously described
(Robertson et al, 2001). All
samples were assayed in duplicate across a single assay. The lower limit of
detection of the immunofluorometric assay was 0.078 ng/mL, with the intraassay
coefficient of variation below 10%.
Inhibin B was measured using a specific ELISA (Oxford BioInnovation, Upper Heyford, United Kingdom) according to the manufacturer's instructions. Rat serum samples were diluted in a dose-dependent manner which was parallel to the standard curve in the assay. The lower limit of detection of inhibin B was 7.8 pg/mL. The within-plate variation was 5.0%.
Concentrations of serum LH were determined using RIA reagents provided by NIDDK (Bethesda, Md). The iodinated standard and antiserum used were rLH I-9 and anti-rLH-S-10, respectively. Rat serum samples were diluted in a dose-dependent manner which was parallel to the standard curve in the assay. The lower limit of detection of rLH was 0.08 ng/mL. The within-assay variation in the LH assay was 10.4%.
Concentrations of testosterone in serum and testicular homogenates were measured without extraction using a direct radioimmunoassay, as described previously (O'Donnell et al, 1994). All samples were assayed in duplicate across a single assay. The lower limit of detection of testosterone was 0.29 ng/mL. The within-assay variation in the testosterone assay was 8.9%.
Statistics
Statistical analysis was carried out using the SigmaStat version 2.1
(Jandel Corporation, San Rafael, Calif) software program. Data were subjected
to a one-way ANOVA, and significance was determined using a Tukey's post hoc
test. In the case of unequal variance, log transformation was carried out to
equal the variance between experimental groups, and significance was
determined using the Student-Newman-Keuls test.
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Results |
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TopAbstractMethods and MaterialsResultsDiscussionReferences |
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Histological Observations
Apoptosis was observed at all stages of the cycle of the seminiferous
epithelium in both control animals and in all LPS-treated groups
(Figure 1a). The location of
these apoptotic cells indicated that they were principally spermatocytes.
Increasing numbers of prematurely released spermatocytes and round spermatids
were observed within the lumen of the seminiferous tubules in the LPS-treated
groups at days 3 and 7 (Figure
1b and c).
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Stereological Analysis
Control testis weights were not significantly different between the
short-term study (1.95 ± 0.02 g/testis; mean ± SEM, n = 7) and
the long-term study (1.90 ± 0.10 g/testis; mean ± SEM, n = 7).
There was no significant effect on testis weight for any treatment group.
In the short-term study, LPS treatment had no significant effect on the number of Sertoli cells, type A spermatogonia, type B spermatogonia, preleptotene spermatocytes, pachytene spermatocytes (at stages IX–XIV), and elongated spermatids (steps 9–14) per testis compared with controls (Table 1). In LPS-treated rats, the numbers of leptotene and zygotene spermatocytes increased by 18% compared with control at day 1, but had returned to normal by day 7 after treatment. Although there was a difference in the number of elongating spermatid (steps 9–14) per testis between day 3 and day 7, neither value was different from the controls. At day 7 after LPS treatment, the number of stage I–VIII pachytene spermatocytes had decreased significantly to 72% of their control values, while the round spermatids had fallen to 77% of the numbers present at day 1.
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In the long-term study, the number of type A spermatogonia, preleptotene spermatocytes, pachytene spermatocytes (stages IX–XIV), elongating spermatids (steps 9–14), and elongated spermatids (steps 15–19) per testis following LPS treatment were the same as control, 28 days after LPS treatment (Table 1). However, there was a significant increase in the number of Sertoli cells (22% increase), type B spermatogonia (33% increase), leptotene and zygotene spermatocytes (21% increase), stages I–VIII pachytene spermatocytes (22% increase), and round spermatids (15% increase) following LPS treatment at day 28. Note that an apparent difference between Sertoli cell numbers between the 2 control groups (approximately 10%) was not statistically significant.
Apoptotic Activity
In the short-term study, the number of apoptotic cells per tubule
cross-section at stages XIV–IV and stages V–VIII showed no change
following LPS treatment compared to control
(Figure 2). On the other hand,
in stages IX–XIII there was an large increase by day 7 following LPS
treatment in the number of apoptotic cells per tubule cross-section (126%
increase), as well as the percentage of tubules containing apoptotic cells
(28.8 ± 3.2% in controls versus 42.9 ± 3.5% in the LPS treated
group at day 7; P < .05). In the long-term study, there was no
significant effect of LPS treatment on the number of apoptotic cells per
tubule cross-section at any stage (Figure
2).
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In the long-term study, intratesticular testosterone in the LPS-treated animals was elevated by more than 2-fold compared with controls, but all other hormones were similar to controls.
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Discussion |
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Remarkably, in spite of the pronounced acute effects on spermatogenesis, this single dose of LPS had only minor consequences for spermatogenic production in the longer term. By 28 days after LPS treatment, one would have expected a significant drop in step 15–19 spermatids, corresponding to the increased loss of pachytene spermatocytes (stages I–VIII) 21 days earlier, and a decline in step 9–14 spermatids due to the increased apoptosis in stages IX–XIII 21 days earlier. Unexpectedly, the numbers of these elongating spermatids were not significantly different from controls. Instead, what was observed was an increase of between 15% and 33% for most other cell types in the seminiferous epithelium, including the Sertoli cells in the LPS-treated testes. The explanation of this result is unclear at present, but it suggests that a compensatory reduction in normal germ cell attrition and possibly an increase in the proliferation rate among some cell types may have occurred, quickly restoring the spermatogenic capacity of the testis to its preinflammation levels. Supporting this suggestion is the observation that there was an apparent trend towards a reduction in the number of apoptotic cells observed at stages XIV–IV and IX–XIII in the long-term LPS-treated group. Moreover, this compensatory activity may have been related to the twofold increase in intratesticular testosterone level in these animals, evidence of increased Leydig cell activity as well. The Leydig cells did not show any obvious morphological alterations, although it is possible that functional changes might be appreciated following more detailed analysis. However, the fact that intratesticular testosterone levels were higher but serum levels of both testosterone and LH were not altered at this time suggests that there may have been an alteration in the testicular vasculature that affects the exit of testosterone from the testis. This also would be consistent with a postinflammatory event.
While increased cell survival and spermatogonial proliferation could account for the compensatory response among the developing germ cells, an apparent increase in Sertoli cell number is harder to explain. It is generally accepted that Sertoli cell number stabilizes in the adult testis, but numbers also decline steadily with age, and seasonal variations may also occur (Johnson, 1986; Dakouane et al, 2005). Although there was an apparent difference between Sertoli cell numbers in the 2 control groups, this was not significant, and the animals were not matched for season or age at time of tissue collection, the 2 experiments being separated by a period of 7 months. Consequently, no conclusion can be drawn as to whether there may have been an age-related or seasonal difference in Sertoli cell number. On the other hand, evidence recently has been obtained from both in vivo and in vitro studies to indicate that adult Sertoli cells can reenter the cell cycle and proliferate following genetic and hormonal manipulation (Chaudhary et al, 2005; Meachem et al, 2005). Altogether, these data suggest that Sertoli cell numbers are relatively stable in the adult, but this involves a low level of continuous turnover, with both loss and recovery of Sertoli cells possible under various conditions. Consequently, the difference in Sertoli cell number in the long-term treatment group could be due to a decrease in normal Sertoli cell loss or stimulation of proliferative activity among these cells. Since IL-1 can act a mitogenic factor for immature Sertoli cells via the p38 MAPK pathway (Petersen et al, 2002; Petersen et al, 2005), this raises the intriguing hypothesis that the up-regulation of inflammatory cytokines during inflammation might stimulate proliferation of adult Sertoli cells, possibly because they also have become disordered in their normal function. Regardless of the explanation, this novel observation that testis hypertrophy/hyperplasia may be a consequence of an acute inflammatory event certainly deserves further investigation. The question also arises: what would happen to spermatogenesis during a more extended period of inflammation?
Although there was no evidence of a significant effect on any hormone parameter following LPS treatment in the present study, at least in the short term, it has been established that LPS-induced inflammation exerts a considerable inhibitory effect on both serum LH and testosterone production in the adult rat within 6 hours after treatment (O'Bryan et al, 2000). Testosterone levels transiently return to normal at the 24-hour time-point, being reduced both before and after this in a biphasic secretion pattern. In this, the rat Leydig cell appears to be quite different in its response to LPS compared with the mouse, which displays a more rapid onset and a more prolonged suppression of testosterone production in particular (Hales et al, 1999). Consequently, the lack of a significant effect on hormones at the day 1 time-point is entirely consistent with previous results, suggesting that the data were obtained during a period of recovery. The fact that all hormone measurements at day 1 appeared to be lower than controls, albeit not significantly lower, does suggest that examination at a slightly earlier period might pick up inhibition of FSH and inhibin B as well, although this is only speculation. In any case, it must be emphasized that the effects of LPS on spermatogenesis observed in the present study were not consistent with withdrawal of either testosterone, which initially affects spermatids at stages VII–VIII, or of FSH, which particularly affects spermatogonia at stages XIV–III (O'Donnell et al, 1994; Meachem et al, 1999). Moreover, testicular testosterone levels cannot be reduced below 30% of control in adult rats with a sublethal concentration of LPS (O'Bryan et al, 2000), and studies have shown that 15%–20% of normal intratesticular testosterone levels are sufficient to maintain full spermatogenesis in the rat (Cunningham and Huckins, 1979; Sharpe et al, 1988). Altogether, the data point towards the involvement of a direct effect on the seminiferous epithelium in LPS-induced inflammation, although the mechanics of those effects remain to be established.
Finally, a less severe testicular phenotype was observed in the present study compared with an earlier study using very similar procedures (O'Bryan et al, 2000). In the earlier study, there was more pronounced epithelial damage with spermatocyte and spermatid loss, spermatogonial apoptosis at stages I–V, and changes in vascular permeability, including microhemorrhage. In the present study, there was no evidence of vasculature disruption or increased apoptosis within stages I–V, and germ cell loss from the seminiferous epithelium appeared to be substantially less. The main reason for the differences between the 2 studies appears to be in severity of the responses to LPS, even though the doses were ostensibly the same. The relatively high mortality rate (25%) observed in the previous study also would seem to support this possibility. It is well recognized that the activity of different preparations of LPS can vary, even when derived from the same microorganism, and the difference in responses may have been related to differences in the potency or composition of the LPS preparations used in the 2 studies. Moreover, there may have been an underlying secondary pathology in the animals used in the previous study. The use of rats maintained under specific pathogen-free conditions in the present study allowed us to avoid this latter complication. Therefore, the present study probably represents a much better indication of the earliest effects of inflammation on spermatogenesis, separate from any confounding external influences. For example, vascular changes could account for the increased spermatogonial apoptosis at stages I–V that was observed only in the earlier study, since these cells are particularly sensitive to an interruption in testicular blood flow (Tjioe and Steinberger, 1970; Turner et al, 1997).
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Acknowledgments |
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Footnotes |
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