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From the * Monash Institute of Reproduction and
Development, Monash University, Melbourne, Victoria, Australia; and
Department of Anatomy and Cell Biology,
Justus-Liebig-University of Giessen, Giessen, Germany.
| Correspondence to: Dr Andreas Meinhardt, Department of Anatomy and Cell Biology, Justus-Liebig-University of Giessen, Aulweg 123, D-35385 Giessen, Germany (e-mail: andreas.meinhardt{at}anatomie.med.uni-giessen.de). |
| Received for publication September 21, 2004; accepted for publication January 4, 2005. |
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Abstract |
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(IFN) are
constitutively expressed by testicular cells, including the Leydig cells. In
the present study, the contribution of the Leydig cell to testicular
inflammatory responses was examined in adult male rats treated with the Leydig
cell–specific toxin, ethane dimethane sulfonate (EDS). Intratesticular
testosterone levels were modulated by subcutaneous testosterone implants.
After 10 days, animals received an injection of lipopolysaccharide (LPS) to
induce an inflammatory response, or saline alone, and were killed 3 hours
later. Both depletion of Leydig cells by EDS and LPS treatment caused a
decrease in collected testicular interstitial fluid to about 35% of control
levels, but the effects were not additive. Maintenance of intratesticular
testosterone reversed the interstitial fluid decline following EDS treatment
and partially prevented the LPS-induced effect. MIF, TGFß1, and
IFN were expressed in both the normal and inflamed testis at similar
levels. In contrast, EDS treatment caused a significant decline in expression
of all 3 cytokines, which was prevented by the testosterone implants. These
data indicate that 1) expression of TGFß1, MIF, and IFN in the
testis is not dependent on the presence of intact Leydig cells but is under
direct testosterone control and 2) the decline in testicular interstitial
fluid during inflammation involves the Leydig cells, acting via both androgens
and nonandrogenic secretions. These data provide further support for a
significant role for the Leydig cell in modulating the testicular response to
inflammation.
Key words: Transforming growth factor-ß, interferon-, macrophage migration inhibitory factor, ethane dimethane sulfonate, testosterone
(TNF), for
example, modulates Leydig cell steroidogenesis
(Xiong and Hales, 1993) and
inhibits germ cell apoptosis by regulating the level of FasL
(Pentikainen et al, 2001), while members of the transforming growth factor-ß (TGFß) family are
implicated in testicular development
(Mullaney and Skinner, 1993;
Teerds and Dorrington, 1993).
Several regulatory cytokines, including macrophage migration inhibitory factor
(MIF) and interferon- (IFN) are produced at significant levels
within the testis even in the absence of inflammation or immune activation
events (Dejucq et al, 1995; Meinhardt et al, 1996). A
better understanding of the production and regulation of these cytokines in
the testis may provide the key to unlocking the processes responsible for
causing testicular damage during inflammation and disease, as well the
maintenance of immune privilege in the testis
(Head and Billingham, 1985; O'Bryan et al, 2000b).
While the local mechanisms that control the expression of cytokines in
normal and diseased testes are poorly understood, evidence points to an
important role for the Leydig cells. There is clear evidence that the Leydig
cells are responsible for recruitment and control of the testicular macrophage
population (Hedger, 2002). Moreover, the Leydig cells and Sertoli cells contribute to the production of
proinflammatory mediators, such as the 2 IL-1 isoforms, IL-6, and inducible NO
synthase (iNOS), during testicular inflammation
(Meinhardt et al, 1996;
Stephan et al, 1997; O'Bryan et al, 2000a;
Soder et al, 2000). Finally,
the immunoregulatory cytokines, TGFß1, MIF, and IFN, are produced
by the Leydig cells either during testicular development or in the adult
(Teerds and Dorrington, 1993;
Dejucq et al, 1995; Meinhardt et al, 1996).
Leydig cells also are responsible for the local regulation, formation, and content of the testicular interstitial fluid (IF), which is of considerable importance to normal testis function (Setchell, 1990). A number of studies have established that IF formation is maintained by androgen-dependent regulation of testicular blood pressure and flow, mediated principally via the seminiferous epithelium (Maddocks and Sharpe, 1989; Collin et al, 1993). Consequently, disruption of the germ cells results in an increase in IF volume, whereas ablation of the Leydig cells by ethane dimethane sulfonate (EDS) causes a decrease in IF volume at least in the short term (Maddocks and Sharpe, 1989; Sharpe et al, 1991). This latter response is quite similar to the effect of lipopolysaccharide (LPS)-induced inflammation on testicular fluid formation, which decreases during inflammation, in direct contrast to the edema response that normally is observed in other tissues (O'Bryan et al, 2000a,b).
It is quite evident, therefore, that the Leydig cell may play an important role in regulating local inflammatory responses in the testis, but direct studies have not yet been performed. Consequently, the following experiments were designed to examine the role of the Leydig cells and testosterone in controlling immunoregulatory cytokine production and IF formation in the normal testis and during inflammation by comparing the responses elicited by the application of the proinflammatory agent lipopolysaccharide (LPS) in the presence and absence of functional Leydig cells in vivo. This model uses a relatively low dose of LPS to induce moderate systemic inflammation in the animals, producing a specific inflammatory response within the testis, which is characterized by a decline in the ability of the Leydig cells to produce testosterone and a unique cascade of inflammatory events, including a transient increase in testicular macrophages without intravasation of neutrophils, attenuated production of the proinflammatory mediators IL-1ß and nitric oxide, and reduction in fluid volume (O'Bryan et al, 2000a,b; Gow et al, 2001; Gerdprasert et al, 2002a,b).
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Materials and Methods |
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Treatment with EDS causes complete destruction of the Leydig cells within 3 days (Kerr et al, 1985; Wang et al, 1994). The transient increase in intratesticular macrophages that accompanies this destruction is back to normal within 10 days, leaving an interstitium that is completely devoid of Leydig cells. In numerous studies it has been established that low-dose subcutaneous testosterone implants (3 cm long; T3 implants) cause a reduction in serum luteinizing hormone (LH) levels, a subsequent decline in intratesticular testosterone, and loss of the developing spermatogenic cells in the testis, while high dose testosterone implants (3 x 8 cm = 24 cm long; T24 implants) similarly cause a reduction in LH but provide sufficient testosterone to maintain intratesticular testosterone levels adequate to support qualitatively normal seminiferous tubule function (Sun et al, 1989; Wang et al, 1994). Rats were separated into the following groups (n = 11 or 12 rats/group): 1) no treatment (controls); 2) T3 subcutaneous implant; 3) dimethyl sulfoxide (DMSO):water (1:3) (1.0 mL/kg) intraperitoneal (DMSO vehicle controls); 4) EDS (75 mg/kg) in DMSO:water intraperitoneal; 5) vehicle plus T24 implant; 6) EDS in DMSO:water plus T24 implant. Ten days later, rats were injected (intraperitoneally) with either endotoxin-free saline or 0.1 mg/kg LPS from Escherichia coli, serotype 0127:B8 (Sigma, Deisenhofen, Germany). Three hours after injection rats were euthanized and tissues were collected. Testis and seminal vesicle weights were determined. One testis was taken for collection of IF as previously described (Wang et al, 1994), and one for isolation of total RNA. Samples of liver were also collected from the animals for controls.
Measurement of Testosterone
Testicular IF was assayed for testosterone without extraction using a
direct 125I-testosterone radioimmunoassay as described previously
(Meinhardt et al, 1998).
RNase Protection Assay for Cytokines
Ribonuclease protection analysis of cytokine expression in rat testes was
performed using a commercially available kit (RiboQuant Multi-Probe, BD
Biosciences, Heidelberg, Germany) according to the manufacturer's
instructions. The custom template set contained the following DNA templates:
TGFß1, IL-6, IL-10, TNF, MIF, and IFN. In addition,
templates for the ribosomal protein L32 and GAPDH housekeeping probes were
included to allow assessment of total RNA levels and normalize sampling. Total
RNA was isolated with the single-step acid phenol-guanidinium
thiocyanate-chloroform extraction method
(Chomczynski, 1993) using
Trizol reagent (Invitrogen, Karlsruhe, Germany). Briefly, 32P
radiolabeled antisense riboprobes were generated by multiplex in vitro
transcription using [-32P]UTP (Amersham, Braunschweig,
Germany) and T7 RNA polymerase. The RNase protection assay was performed
according to the manufacturer's protocol. Radiolabeled probes were hybridized
with 5 µg of total testicular or liver RNA samples at 56°C overnight.
After RNase A/T1 treatment for digestion of single-stranded RNA, samples
together with undigested probes and in vitro transcribed 100 bp RNA markers
(Century Marker, Ambion, Huntingdon, UK) were resolved on a 50 x 30 cm
5%–6% polyacryamide/7M urea gel
(Table). The gel was dried for
1 hour at 80°C under vacuum (Bio-Rad, München, Germany), exposed to a
phosphorimage screen overnight, and analyzed on a Fuji PhosphorImager FLA3000G
(Raytest, Straubenhardt, Germany). Subsequently, RNase protected bands were
quantified using Fuji Image Gauge software. For each treatment group analyzed,
RNA was prepared from 4 randomly selected animals. The probe set was labeled 3
to 4 separate times for analyses of samples from each testis examined. Liver
RNA (±LPS) was included as control for cytokine expression. All
cytokine data were normalized for variation in loading against L32 and GAPDH.
Both normalizations yielded very similar results, but only the L32 normalized
data are presented.
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Statistics
Data were analyzed by 2-way analysis of variance (ANOVA) after appropriate
transformation to normalize the data and equalize variance, in conjunction
with the Student Newman-Keuls multiple-range test, using SigmaStat version 1.0
software (Jandel Scientific Software, San Rafael, Calif). Statistical
comparisons were limited to groups receiving the same injection vehicle, that
is, either saline alone or DMSO:water and saline, to avoid complications due
to the minor effects of DMSO on testicular IF volume and permeability
characteristics (Hedger and Hettiarachchi,
1994; Wang et al,
1994). Comparisons between experimental groups were considered
statistically significant at the P < .05 level.
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Results |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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IF Volume Regulation
Neither the T3 nor T24 implants on their own had a significant effect on
the volume of recovered IF, but EDS caused a 65% decline in volume, which was
prevented by the T24 implants (Figure
2). Treatment with LPS caused a decrease of about 70% in IF volume
in control rats by 3 hours after treatment
(Figure 2). In contrast to the
suppression by EDS, which led to a consistent reduction of between 31% and 56%
(range: 27–49 µL) compared with controls, the variation in the degree
of suppression by LPS was very large (1–86 µL). In fact, the response
to LPS did not actually achieve significance in the T3-implanted animals due
to the variation in IF volumes and a slight, but not statistically
significant, reduction in IF volume of the corresponding saline-injected
controls (Figure 2). However,
LPS clearly had no effect on IF volume in EDS-treated rats, and the effects of
LPS on IF volume were at least partially prevented in the T24 implanted rats
compared with controls (Figure
2).
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From each experimental group 3 to 4 samples were randomly selected for
RNase protection assay (Figure
3). Signals were detected for TGFß1 and MIF in all testes
samples. Expression levels for IL-6, IL-10, TNF-, and IFN were
either very weak or not detectable even in the LPS treatment groups and
therefore not quantifiable (Figure
3). In contrast, all cytokines examined were detected either in
liver (not shown) or in the control RNA provided by the kit
(Figure 3). T3 implants caused
a slight but significant increase in MIF and IFN expression, but not in
TGFß1 mRNA levels compared with untreated control
(Figure 4A through C). LPS had
no significant effect on these cytokines in control and T3 testes. EDS,
however, caused a moderate reduction in MIF expression and a large decrease in
both TGFß1 and IFN expression
(Figure 4A through C). LPS
potentiated this effect of EDS on MIF, TGFß1, and IFN mRNA levels,
while T24 implants prevented the decrease. IFN also revealed a slight
increase in T24 implanted rats (Figure
4C).
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Discussion |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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The volume of IF collected from the rat testis overnight via an incision in the testicular capsule is directly proportional to the volume of fluid present in the extratubular space, with only minor contributions from the other testicular compartments (Setchell and Sharpe, 1981; Sharpe and Cooper, 1983). As has been observed previously, treatment with EDS caused a consistent 60% to 70% fall in the volume of IF recovered (Sharpe et al, 1990; Hedger et al, 1998). LPS treatment alone caused a similar average fall in IF volume (O'Bryan et al, 2000b), but it should be noted that LPS induced a much more variable suppression of IF volumes than did EDS treatment, at least within the first 3 hours after LPS injection. This response is in contrast to that of other tissues where inflammation normally causes an increase in blood flow and vascular permeability, followed by edema. That there is a local inflammatory response in the testis following treatment with LPS at this time point has been established by the fact that several proinflammatory mediators, including IL-1ß, monocyte chemoattractant protein-1, and iNOS, are up-regulated (O'Bryan et al, 2000a; Gow et al, 2001; Gerdprasert et al, 2002b) and there is neutrophil accumulation in the testicular blood vessels and a small increase in endothelial cell leakage in spite of the overall fall in IF volume (O'Bryan et al, 2000b). The large variability in the suppression of fluid volume from animal to animal is consistent with our previous observations (O'Bryan et al, 2000b) and suggests that this is a characteristic feature of the inflammatory response in the rat testis. In the EDS-treated testis, the mean and variability of IF volumes were almost identical in saline-treated and LPS-treated testes, indicating that in the absence of the Leydig cells, LPS was no longer able to cause the very large suppression in IF volume responses that was observed in some control animals.
The effects of LPS also appeared to be partially, but not completely, reduced in testes of rats in which testicular androgen levels are maintained at a level that supports close to normal spermatogenesis by the use of T24 implants. A slightly different result was observed in the T3-implanted animals, with a minor, but not significant, reduction in pretreatment IF volume, which might be attributable to the reduced intratesticular testosterone levels. Although LH has stimulatory effects on IF volume in vivo and there is some evidence that LH binds to receptors on the testicular endothelium (Veijola and Rajaniemi, 1986; Bergh et al, 1990; Ghinea et al, 1994), the changes in IF volume observed in the present study were not consistent with any significant direct role for this hormone; that is, IF volume was largely unaltered by T implants that completely suppress circulating LH levels, while EDS treatment reduced IF volume even though LH levels would be dramatically elevated in these animals. Altogether, these data indicate that the inhibitory effect of LPS on IF volume is dependent on the presence of intact Leydig cells, and although changes in intratesticular androgen levels influence this response, other products of the Leydig cells also appear to be involved. This response may involve disruption of the putative Leydig cell–mediated vascular regulatory agent(s) (Veijola and Rajaniemi, 1986; Damber et al, 1992; Collin and Bergh, 1996), which has not yet been identified. Physiological responses that could cause a decline in testicular IF volume include an increase in intratesticular pressure from swelling of the seminiferous tubules or contraction of the capsule, increased lymph flow or venous resorption, or a reduction in blood pressure or flow (Setchell et al, 1994). It is possible that the up-regulation of iNOS and increased local production of the vasodilator NO, leading to reduced blood pressure and flow in the testis, may be involved, since the Leydig cells are the earliest and most effective producers of iNOS in the LPS-treated rat testis (O'Bryan et al, 2000a). Whatever the mechanism, the data clearly indicate that in spite of increases in vascular permeability in the inflamed testis (O'Bryan et al, 2000b), which in other tissues result in edema, the inflammatory response of the testis causes additional vascular changes leading to a range of vascular responses, but generally producing an overall reduction in the normally large volume of testicular IF.
As expected, LPS had no significant effect on testis or seminal vesicle weights over the short time frame of the treatment (3 hours). LPS appeared to have no significant effect on IF testosterone levels in the control groups over this time frame as well. This lack of a significant effect on IF testosterone at this time is consistent with a previous observation that, while Leydig cell testosterone production is obviously affected much earlier, maximal reduction of testosterone levels in the testicular IF does not occur until 6 hours after treatment with LPS in the rat testis (O'Bryan et al, 2000b). It should be noted that there are distinct species differences in the dynamics of inhibition of steroidogenesis by LPS, since Leydig cell inhibition responses in mice generally have a more rapid onset and are larger and more prolonged than those of rats (Bosmann et al, 1996; Sewer and Morgan, 1998; Hales et al, 2000; O'Bryan et al, 2000b; Gow et al, 2001).
With a ribonuclease protection assay, expression of TGFß1, IFN,
and MIF was clearly demonstrated in the testis, whereas IL-6, TNF, and
IL-10 mRNAs were barely detectable using this approach, in spite of the fact
that the production of at least IL-6 and TNF- has been described in the
adult testis (De et al, 1993; Syed et al, 1993). However,
previous studies either used extremely sensitive methods such as reverse
transcription–polymerase chain reaction or investigated synthesis and
production in isolated cells or a combination of both rather than total testis
(De et al, 1993;
Xiong and Hales, 1993;
Kern et al, 1995;
Huang et al, 2003), suggesting
that the sensitivity of the assay was insufficient to detect expression of low
copy number cytokines in total testes RNA. Conversely, this also suggests that
TGFß1, IFN, and MIF are relatively highly expressed cytokines in
the testis.
Ablation of the Leydig cells by EDS treatment did not completely eliminate
production of TGFß1, IFN, or MIF before inflammation, indicating
that this cell type is not required for their expression in the testis.
However, expression of all cytokines investigated was significantly reduced by
EDS treatment and restored by T24 implants, strongly suggesting that their
expression is androgen regulated. Both these observations implicate the
Sertoli cells and peritubular cells as the main source of these cytokines in
the Leydig cell–depleted testis
(Teerds and Dorrington, 1993;
Dejucq et al, 1995;
Meinhardt et al, 1999). The
fact that the reduced intratesticular androgen levels caused by T3 implants
did not cause a similar decline in the expression of these cytokines suggested
that the level of testosterone required may actually be quite low or that the
extent of the damage to spermatogenesis may be an important determinant as
well. In fact, 2 of the cytokines (MIF, IFN) were elevated in
T3-implanted animals, suggesting that testosterone may have a biphasic effect
on their expression. Inflammation did not increase the expression of MIF,
TGFß1, and IFN mRNA. On the contrary, an additive down-regulatory
effect of LPS for these cytokines was observed in the EDS-treated testes. The
mechanism by which LPS further inhibits expression of these cytokines in the
EDS-treated testis remains to be determined. It appears unlikely that a direct
effect of LPS on the Sertoli or peritubular cells acting via the LPS receptor
signaling pathway is involved, and it is possible that the reduction is due to
an immunomodulatory intermediate produced by the testicular macrophages.
Overall, the data suggest that the complete lack of androgens suppresses
production of these cytokines. However, treatment with T3 implants showed that
only very low levels of intratesticular testosterone are required to maintain
normal expression levels, clearly indicating that it is testosterone, and not
an indirect effect of androgens mediated via the seminiferous epithelium, that
is responsible for this maintenance. Moreover, the failure of these cytokines
to display acute regulation by inflammation in the testis may be related both
to the fact that they are produced constitutively by testicular cells that are
relatively insensitive to LPS stimulation (ie, testicular somatic cells and
germ cells) and the unique regulatory phenotype of the testicular macrophage
(Hedger, 2002).
These data also confirm that Leydig cells are not the sole source of any of
the above cytokines in the testis. IFN has been localized to Sertoli
cells and postmeiotic germ cells in addition to Leydig cells
(Dejucq et al, 1995). The 3
mammalian TGFß isoforms (1–3) are very highly expressed by Sertoli
cells, peritubular cells, and Leydig cells in the fetal and immature testis,
although production declines dramatically postpuberty
(Mullaney and Skinner, 1993;
Avallet et al, 1994;
Konrad et al, 2000). In the
postpubertal testis, they have been localized to the developing germ cells in
a developmentally specific pattern of expression
(Teerds and Dorrington, 1993;
Caussanel et al, 1997). In
addition, immature germ cells secrete bioactive TGFß
(Haagmans et al, 2003).
Although MIF has been localized exclusively to the Leydig cells in the normal
adult rat testis (Meinhardt et al,
1996), as noted previously, MIF production is not lost after
depletion of the Leydig cells because compensatory MIF production by Sertoli
cells occurs in the Leydig cell–depleted testis
(Meinhardt et al, 1999). It is
possible that similar compensatory production may occur for other cytokines in
the testis. Finally, activation of the testicular macrophages or recruitment
of leukocytes in the EDS- and LPS-treated testis could also influence the
pattern of production that was observed
(Wang et al, 1994). The data
also suggest that, although a role in maintaining IF volume in the normal
testis cannot be excluded, none of the 3 cytokines appears to be responsible
for the changes in IF volume during LPS-induced inflammation.
Taken together, these data are consistent with a direct effect of androgen on the expression of several cytokines in the testis that are produced at a high constitutive level. The pattern of expression for these cytokines (ie, apparently high level of endogenous production, stimulation by androgens or spermatogenic cells, absent or limited response to inflammation) is unique to the testis and may have important implications for the maintenance of the unique immune environment of the testis.
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Acknowledgments |
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Footnotes |
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) or 0.1 mg/kg LPS intraperitoneally
(
)—experimental groups were normal controls (control),
DMSO-treated controls (DMSO), EDS treatment (EDS), 3-cm testosterone implants
(T3), and 24-cm testosterone implants (T24) in combination with DMSO or EDS.
(A) testis weight; (B) seminal vesicle weight; and (C) IF
testosterone. All values are mean ± SEM of 5–6 animals.
Statistical comparisons were limited to groups receiving the same injection
vehicle, either saline alone (left-hand graph) or DMSO:water and saline
(right-hand graphs). Experimental group values with same letter superscript
are not significantly different (P > .05). Significant changes in
organ weights are a reflection of circulating testosterone concentrations.
There were no significant differences between values for the saline-injected
and LPS-treated testes in any experimental group.
