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Endocrinology Vol. 138, No. 9 3719-3726
Copyright © 1997 by The Endocrine Society

ARTICLES

Decreased Cyclin A2 and Increased Cyclin G1 Levels Coincide with Loss of Proliferative Capacity in Rat Leydig Cells During Pubertal Development1

Ren-shan Ge and Matthew P. Hardy

The Population Council (R.-S.G., M.P.H) and Rockefeller University (M.P.H.), 1230 York Avenue, New York, New York 10021

Address all correspondence and requests for reprints to: Matthew P. Hardy, The Population Council, 1230 York Avenue, New York, New York 10021. E-mail: hardy{at}popcbr.rockefeller.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Postnatal development of Leydig cells can be divided into three distinct stages of differentiation: initially they exist as mesenchymal-like progenitors (PLC) by day 21; subsequently, as immature Leydig cells (ILC) by day 35, they acquire steroidogenic organelle structure and enzyme activities but metabolize most of the testosterone they produce; finally, as adult Leydig cells (ALC) by day 90 they actively produce testosterone. The aims of the present study were to determine whether changes in proliferative capacity are associated with progressive differentiation of Leydig cells, and if the proliferative capacity of Leydig cells is controlled by known hormonal regulators of testosterone biosynthesis: LH, insulin-like growth factor I (IGF-I), androgen, and estradiol (E2). Isolated PLC, ILC, and ALC were cultured in DMEM/F-12 for 24 h followed by an additional 24 h in the presence of LH (1 ng/ml), IGF-I (70 ng/ml), 7{alpha}-methyl-19-nortestosterone (MENT, 50 nM), a synthetic androgen that is not metabolized by 5{alpha}-reductase, or E2 (50 nM).

Proliferative capacity was measured by assaying [3H]thymidine incorporation and labeling index (LI). Messenger RNA (mRNA) and protein levels for cyclin A2 and G1, which are putative intracellular regulators of Leydig cell proliferation and differentiation, were measured by RT-PCR and immunoblotting, respectively. Thymidine incorporation was highest in PLC (9.24 ± 0.21 cpm/103 cell, mean ± SE), intermediate in ILC (1.74 ± 0.07) and lowest in ALC (0.24 ± 0.03). Similarly, LI was highest in PLC (13.42 ± 0.30%, mean ± SE), intermediate in ILC (1.95 ± 0.08%), and undetectable in ALC. Cyclin A2 mRNA levels, normalized to ribosomal protein S16 (RPS16), were highest in PLC (2.76 ± 0.21, mean ± SE), intermediate in ILC (1.79 ± 0.14), and lowest in ALC (0.40 ± 0.06). In contrast, cyclin G1 mRNA levels were highest in ALC (1.32 ± 0.16), intermediate in ILC (0.47 ± 0.07), and lowest in PLC (0.12 ± 0.02). The relative protein levels of cyclin A2 and G1 paralleled their mRNA levels. Increased proliferative capacity was observed in PLC and ILC, but not ALC, after treatment with either LH or IGF-I. Treatment with MENT increased proliferative capacity only in ILC and had no effect in any other group. Treatment with E2 decreased proliferative capacity in PLC but not in ILC or ALC. The changes in proliferative capacity after hormonal treatment paralleled cyclin A2 mRNA and were the inverse of cyclin G1 mRNA levels. We conclude that: 1) decreased cyclin A2 and increased cyclin G1 are associated with the withdrawal of the Leydig cell from the cell cycle; 2) the proliferative capacity of Leydig cells is regulated differentially by hormones and is progressively lost during postnatal differentiation.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE RATE OF testosterone production depends on both the steroidogenic capacity of individual Leydig cells and their total number per testis. The total number of Leydig cells is determined during pubertal development, because their proliferative activity is limited to the prepubertal period (reviewed in 1 . Hormonal control of Leydig cell proliferation is poorly understood, but several factors are involved, including LH, insulin-like growth factor-I (IGF-I), androgen, and estrogen. Administration of LH or its analogue, human CG, to adult rats increases Leydig cell numbers (2, 3). However, it is still unclear in the immature rat whether LH stimulates Leydig cell division or the division of Leydig cell precursors. IGF-I stimulates mitosis in many tissues (reviewed in 4 . Receptors for IGF-I are present in Leydig cells and their precursor cells (5), and the concentration of IGF-I in interstitial fluid is highest during puberty (6). This suggests that IGF-I facilitates Leydig cell development, possibly by increasing proliferation. Androgen may also induce proliferation of Leydig cells because androgen insensitivity in tfm, testicularly feminized mice, leads to decreased Leydig cell number (7, 8). In contrast to LH, IGF-I, and androgen, estrogen is known to inhibit Leydig cell development because administration of estrogen to neonatal rats significantly reduces Leydig cell numbers in adults (9, 10). This suggests that Leydig cell proliferation is also affected directly and/or indirectly by this steroid.

As in all cells, Leydig cell proliferation is postulated to be controlled by protein complexes composed of a cyclin, the regulatory subunit, and a cyclin-dependent kinase, the catalytic subunit (reviewed in 11 . Regular occurrence of cell division is mediated by differential synthesis and degradation of cyclins at specific points during the cell cycle (12). Nine different classes of vertebrate cyclins have been identified to date, designated A through I (12, 13, 14, 15). Cyclin A is a known mediator of cell proliferation (16, 17, 18), and two forms, A1 and A2, are present in mouse testis (19, 20). Localization of messenger RNA (mRNA) of the two forms in the mouse testis by hybridization histochemistry indicates that cyclin A1 stimulates germ cell meiosis (20), whereas cyclin A2 controls mitosis in both somatic cells and germ cells (16, 17, 18, 20). In contrast to all eight other cyclin classes, cyclin G lacks a "destruction box" amino acid sequence necessary for degradation, and contains an epidermal growth factor receptor-like autophosphorylation motif (21). Two types, cyclin G1 and G2, have been detected in human tissues (22). The cyclin G1 gene is transcriptionally activated by the p53 tumor suppressor protein (23, 24, 25, 26), indicating that, unlike the other eight cyclin classes, G1 participates in control of cell growth, differentiation and/or apoptosis.

Given the disparate roles of cyclins A2 and G1 in other cell types, the initial aim of the present study was to determine whether steady state levels of cyclin A2 and G1 mRNAs vary with Leydig cell proliferative capacity during development. Postnatal development of Leydig cells can be divided, conceptually, into three distinct stages of differentiation: initially they exist as mesenchymal-like progenitors (PLC) by day 21; subsequently, as immature Leydig cells (ILC) by day 35, they acquire steroidogenic organelle structure and enzyme activities but metabolize most of the testosterone they produce; finally, as adult Leydig cells (ALC) by day 90 they actively produce testosterone (reviewed in 1 . The proliferative capacities of these three distinct stages of pubertal differentiation were evaluated with respect to their sensitivities to hormonal regulation. This analysis showed that declining Leydig cell proliferative capacity during postnatal development was associated with decreased cyclin A2 and increased cyclin G1 mRNAs. Modulation of proliferative capacity and cyclin A2 and G1 mRNA levels indicated that hormonal regulators acted on Leydig cells in a stage specific manner.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals
Sprague-Dawley rats (dams with litters of male pups, immature males, and adult males) were purchased from Charles River Laboratories (Wilmington, MA). Male rats were 21, 35, and 90 days of age on the day of Leydig cell isolation. The animals were killed by asphyxiation with CO2. The animal protocol was approved by the Institutional Animal Care and Use Committee of the Rockefeller University (Protocol 91200).

Cell isolation
A complete description of the cell isolation procedure has been published (27, 28). In brief, testes from 40 21-day-old rats were removed for isolation of PLC. Decapsulated testes were dispersed with 0.25 mg/ml collagenase (collagenase-D, Boehringer Mannheim Biochemicals, Indianapolis, IN) in medium 199 for 10 min at 34 C with shaking. The separated cells were filtered through two layers of nylon mesh, centrifuged at 250 x g, and resuspended in 55% isotonic Percoll. Following density gradient centrifugation at 25,000 x g for 45 min at 4 C, the PLC fraction was collected between densities of 1.064 and 1.070 g/ml. The cells were washed with HBSS, centrifuged at 250 x g, and resuspended in phenol red-free medium (DMEM-Ham’s F-12, D-2906, Sigma Chemical Co., St. Louis, MO) supplemented with 1 mg/ml BSA.

ILC were isolated from the testes of 20 35-day-old rats, with the following modifications to the above procedure. Testes were perfused with 1 mg/ml collagenase in medium 199 via the testicular artery before decapsulation. The ILC fraction was collected from the Percoll gradient between densities of 1.07 and 1.088 g/ml. ALC were purified from the testes of 6 90-day-old rats according to the method of Klinefelter et al. (28). Before the Percoll density gradient centrifugation, collagenase-dispersed interstitial cells were elutriated in the Beckman JE-6B elutriation chamber (Palo Alto, CA) at a flow rate of 16 ml/min at 2,000 rpm, after which ALC were collected from the Percoll gradient between densities of 1.07 and 1.09 g/ml. Purities of Leydig cell fractions were evaluated by histochemical staining for 3ß-hydroxysteroid dehydrogenase activity, with 0.4 mM etiocholanolone as the steroid substrate (29). Enrichment of PLC was typically to 90% purity (approximately 90% of the cells were lightly stained). Of the remaining 10%, 6% were intensely stained. Based on previous cytological results (30), of the 4% that were unstained, less than 1% were macrophages. ILC and ALC were typically enriched to 92–95% and were stained intensely.

Cell culture
Leydig cells were cultured for 48 h in phenol red-free medium (DMEM-Ham’s F12) supplemented with 1 mg/ml BSA, 1 mg/ml bovine lipoprotein, and 25 mM HEPES (pH 7.2) in a 34 C, 5% O2, 5% CO2 humidified incubator. In groups that received hormonal treatment, Leydig cells were first cultured for 24 h in hormone-free medium. The media were then removed, and replaced with fresh medium containing either: 1 ng/ml ovine LH (a gift from NIDDK); 70 ng/ml of IGF-I (Mallinckrodt, Chesterfield, MO); 50 nM 7{alpha}-methyl-19-nortestosterone (MENT, kindly provided by The Upjohn Company, Kalamazoo, MI); or 50 nM E2 (Sigma) for the final 24 h in vitro. Because immature stages of the Leydig cell contain high levels of the androgen metabolizing enzymes, 5{alpha}-reductase and 3{alpha}-hydroxysteroid dehydrogenase (30, 31), MENT, which is not metabolized by 5{alpha}-reductase (32), was used in the present study to examine the androgen action (33).

Thymidine incorporation
Leydig cells were labeled with [3H]thymidine (DuPont-New England Nuclear, Boston, MA) at 1 µCi/ml (specific activity 104.7 Ci/mmol) during the last 2 h of incubation. After labeling, the cells were washed twice with Dulbecco’s PBS and harvested. Cells were lysed in 0.5 ml hyamine hydroxide (ICN Radiochemicals, Irvine, CA) and radioactivity was measured in a liquid scintillation counter.

Autoradiographic determination of Leydig cell labeling index
Leydig cells were also grown in 8-chamber culture slides (Lab-Tek, Nunc, Naperville, IL), hormonally treated in vitro, and labeled with [3H]thymidine as described above. Cells were then washed twice with Dulbecco’s PBS, fixed in 4% formaldehyde, and stored in 70% ethanol until autoradiography. Developed silver grains associated with radiolabeled cells were observed by light microscopy at a magnification of 160x. Cells having more than five silver grains overlying the nucleus were judged to be labeled. A total of 200 cells in each culture well was counted, and the labeling index (LI) were calculated as the number of labeled cells/total number of cells counted x 100.

RT-PCR analysis of cyclin A2 and G1 mRNA levels
Rat cyclin A2 and G1 mRNA levels were determined by RT-PCR. Total RNA was extracted from isolated Leydig cells by a single-step method, using phenol and guanidinium thiocyanate (Trireagent, Molecular Research Center, Inc., Cincinnati, OH) according to the manufacturer’s instructions. Leydig cell total RNA (400 ng) was reverse transcribed with avian myeloblastosis virus reverse transcriptase (Promega) in the presence of random hexamer plus dNTPs at 42 C for 75 min, and the reaction was terminated by heating at 95 C for 5 min. Cyclin A2, or cyclin G1, complementary DNA (cDNA) sequences were coamplified with endogenous ribosomal protein S16 (RPS16) cDNA sequence as an internal standard. PCR was initiated by Taq DNA polymerase in the presence of [{alpha}-32 P] dCTP and proceeded for 30 cycles at an annealing temperature of 50 C. Because the cDNA sequences of cyclin A1 and A2 in rat have not been cloned, the primer sequences for cyclin A2 (forward 5'-CGTGGACTGGTTAGTTGA-3'; reverse 5'-ATGGCAAATACTTGAGGT-3') were based on the published human and mouse cyclin A2 cDNA sequences (20, 34). The primer sequences of cyclin G1 (forward 5'-CCTTCCAATTTCTGCAGCTC-3'; reverse 5'-CTTGGAAACAAGCTCTTGCC-3') were based on the published rat cyclin G1 cDNA sequences (21). PCR products of cyclin A2 (0.42 kb) and cyclin G1 (0.28 kb) were sequenced by the protein/DNA Technology Center at Rockefeller University. Using the LFASTA sequence analysis program (35), the rat cyclin A2 partial sequence was checked against corresponding regions in mouse cyclin A1 (GenBank accession number X84311), human and mouse cyclin A2 (GenBank accession numbers X51688 and X75483). The partial cyclin G1 sequence was checked against rat cyclin G1 (GenBank accession number X70871). The sequence of the PCR product (0.15 kb) formed using the internal control RPS16 primers was confirmed by restriction analysis according to Shan et al. (36). Radiolabeled PCR bands were visualized on Kodak imaging film (Eastman-Kodak, Rochester, NY). Quantitative analysis of mRNA levels was performed by scanning films in a densitometer (LKB Ultroscan, Bromma, Sweden). The signal intensities for cyclin A2 and G1 mRNAs were normalized to RPS16 and the measurements are expressed in arbitrary units.

Western blotting analysis of cyclin A and cyclin G
Leydig cells were homogenized and boiled in equal volumes of sample loading buffer. Homogenized samples (50 µg protein) of PLC, ILC, and ALC were electrophoresed on a 10% SDS polyacrylamide gel (37). Proteins were electroblotted onto a nitrocellulose membrane, and the membranes were incubated with a rabbit polyclonal antihuman cyclin A antibody (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) or a rabbit polyclonal anti-rat cyclin G antibody diluted to 1:5000, after having been blocked with a 10% aqueous solution of nonfat dry milk powder. The membranes were then washed and incubated with a 1:5000 dilution of goat antirabbit antiserum that was conjugated to horseradish peroxidase (Amersham, Arlington Heights, IL). The washing step was repeated, and immunoreactive bands were visualized by chemiluminescence according to the manufacturer’s protocol. Protein levels were measured by densitometry of the films, and ILC and ALC were calculated as percentages of PLC signal intensity.

Data analysis
All measurements were repeated at least three times. The data were analyzed by the Kruskal-Wallis ANOVA followed by Fisher’s LSD method of multiple comparisons testing to identify significant differences between group (38). Differences were regarded as statistically significant at P < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Proliferative capacity of Leydig cells
Developmental trends of proliferative capacity in Leydig cells were first characterized by measuring DNA synthesis at three stages of differentiation, PLC on day 21, ILC on day 35, and ALC on day 90. As shown in Fig. 1aGo, the highest rate of [3H]thymidine incorporation occurred in PLC (9.24 ± 0.21 cpm/103 cells, mean ± SE). Labeling was intermediate in ILC (1.74 ± 0.07 cpm/103 cells) and negligible in ALC (0.24 ± 0.03 cpm/103 cells). The LI was measured to estimate the proportion of Leydig cells that were mitotically active, was highest in PLC (13.42 ± 0.30%, mean ± SE), intermediate in ILC (1.95 ± 0.08%), and was undetectable in ALC (Fig. 1bGo). These results indicate that Leydig cells lose their proliferative capacity during postnatal development, in agreement with previous studies (39, 40).



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  		Figure 1. Developmental changes in Leydig cell proliferative capacity. PLC, ILC, and ALC were isolated and cultured as described in Materials and Methods. Radioactive thymidine incorporation during the last 2 h in vitro was measured by scintillation counting (a). Values represent means ± SE for nine assays from three separate experiments. Cells undergoing nuclear incorporation of [3H] thymidine were assessed by for LI after autoradiography (b). Values represent means ± SE for six wells from three separate experiments. Shared alphabet letters indicate when there was not a difference at P < 0.05.

 
Cyclin A2 and G1 mRNA levels in Leydig cells
Cyclins were evaluated because they are hypothesized to be critical regulators of the Leydig cell cycle. Cyclin A2 provided an index of increased proliferative capacity, whereas cyclin G1 was an index for terminal differentiation or apoptosis (17, 18, 19). The rat cyclin A2 cDNA sequence has not been established. A pair of primers containing a sequence shared by human and mouse cyclin A2 cDNA (20, 34) was used to detect a partial sequence of cyclin A2 in rat Leydig cells. After RT-PCR, a unique PCR product (0.42 kb) was observed in samples of rat Leydig cell RNA (Fig. 2aGo). The sequence for the PCR product had 90% similarity with human cyclin A2, 96% similarity with mouse cyclin A2, but only 67% similarity with mouse cyclin A1. This indicated that cyclin A2 mRNA was present in rat Leydig cells. The steady-state levels of cyclin A2 mRNA were measured by RT-PCR with coamplification of RPS16 mRNA sequence as an internal control. Quantitative estimates of mRNA level were obtained using RT-PCR assays performed under conditions generating a linear range of specific amplification of cyclin A2, G1 and RPS16 mRNAs. The relationship between cycle number and RT-PCR amplification of cyclin A2, G1, and RPS16 mRNAs starting from a fixed concentration of Leydig cell RNA (1/5 of cDNA generated from 2 µg RNA) was determined and was found to be linear for up to 35 cycles (data not shown). Subsequent measurements were performed using 30 cycles of amplification. After normalization with RPS16 mRNA (Fig. 2bGo), PLC had the highest level of cyclin A2 mRNA (2.76 ± 0.21, mean ± SE), ILC were intermediate (1.79 ± 0.14) and ALC were lowest (0.40 ± 0.06). These results suggested that cyclin A2 mRNA decreased during Leydig cell differentiation.



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  		Figure 2. Developmental changes in cyclin A2 and G1 mRNA levels in Leydig cells. Total cellular RNA was extracted from PLC, ILC, and ALC after 48 h culture in hormone-free medium. A 0.42-kb PCR fragment was detected using primers based on human and mouse cyclin A2 cDNA sequences (a). Steady-state levels of cyclin A2 mRNA were measured by RT-PCR after coamplification with RPS16, an internal control (b). A 0.28-kb PCR fragment was observed in Leydig cells and was identified as cyclin G1 (c). Steady-state levels of cyclin G1 mRNA were measured by RT-PCR after coamplification with RPS16 (d). Values normalized to RSP16 represent means ± SE for three to five separate experiments. Shared alphabet letters indicate when there was not a difference at P < 0.05.

 
Cyclin G1 mRNA was detected using primers based on the published sequence for rat cyclin G1 mRNA (21, 22). A 0.28-kb PCR product was observed in Leydig cells (Fig. 2cGo), and its authenticity was confirmed by sequencing. Analysis of steady state mRNA levels showed that, in contrast to cyclin A2, cyclin G1 mRNA levels were lowest in PLC (0.12 ± 0.02, mean ± SE), intermediate in ILC (0.47 ± 0.07), and highest in ALC (1.32 ± 0.16) (Fig. 2dGo). This indicated that cyclin G1 mRNA increased during Leydig cell differentiation.

Protein levels of cyclin A and G in Leydig cells
Cyclin levels were analyzed further by immunoblotting samples of purified Leydig cells for cyclin A2 and G1 proteins. The anticyclin A antiserum recognized a 56-kDa cyclin A2 protein in rat Leydig cells (Fig. 3aGo), consistent with the known molecular mass of cyclin A2. Relative cyclin A2 protein levels changed in parallel with its steady state mRNA described above. Cyclin A2 protein in ILC and ALC was 67.1 ± 4.4% (mean ± SE) and 15.1 ± 5.1% of PLC, respectively (Fig. 3bGo). The anticyclin G antiserum recognized a 32 kDa cyclin G1 protein in rat Leydig cells (Fig. 3cGo), consistent with the known molecular mass of cyclin G1. Cyclin G1 protein also changed in parallel with its mRNA. Protein levels in ILC and ALC were 318.6 ± 13.0% and 827.4 ± 134.2% of PLC, respectively (Fig. 3dGo). These results indicate that cyclin A2 decreased, and cyclin G1 increased, during Leydig cell differentiation.



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  		Figure 3. Western blot analysis of Leydig cell cyclin A2 and G1 proteins in Leydig cells. The homogenates of PLC, ILC, and ALC were electrophoresed and transferred to membrane for detection the 56 kDa cyclin A2 protein (a) and 32 kDa cyclin G1 protein (b). Quantification of cyclin A2 and G1 immunoreactivity was performed by scanning densitometry. Relative protein levels of cyclin A2 (c) and G1 (d) in ILC and ALC were normalized to PLC. Values represent means ± SE for three separate experiments. Shared alphabet letters indicate when there was not a difference at P < 0.05.

 
Effects of hormonal treatment on Leydig cell proliferative capacity and cyclin A2 and G1 mRNA levels
Changes in the proliferative capacity of Leydig cells during pubertal development might require differential sensitivity of their intermediate precursors to hormonal regulation. Therefore, Leydig cells were cultured in the presence of known modulators of their structure and steroidogenic function: LH (1 ng/ml), IGF-I (70 ng/ml), MENT (50 nM), and E2 (50 nM). As shown in Fig. 4Go, both LH and IGF-I increased proliferative capacity in PLC and ILC, but had no effect on ALC. Treatment with the androgen, MENT, increased proliferative capacity only in ILC (2.88 ± 0.24 vs. 1.74 ± 0.07 cpm/103 cell in the control) and had no effect on PLC or ALC. Treatment with E2 decreased proliferative capacity in PLC (7.53 ± 0.26 vs. 9.24 ± 0.21 cpm/103 cell in the control) but had no effect on ILC or ALC. These results indicated that distinct sets of hormonal factors regulate proliferation in Leydig cells during pubertal differentiation.



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  		Figure 4. Effects of hormonal treatment on proliferative capacity in Leydig cells. PLC, ILC, and ALC were isolated from 21-, 35-, and 90-day-old rats, respectively, and cultured in DMEM/F-12 with 1 mg/ml BSA and 1 mg/ml bovine lipoprotein for 24 h, then for another 24 h in the absence (control) or presence of the following hormones: LH (1 ng/ml), IGF-I (70 ng/ml), MENT (50 nM) and E2 (50 nM). Radioactive thymidine incorporation during the last 2 h in vitro was measured by scintillation counting (a). Values represent means ± SE for nine assays from three separate experiments. Cells undergoing nuclear incorporation of [3H]thymidine were assessed by for LI after autoradiography (b). ND represents undetectable signal. Values represent means ± SE for six wells from three separate experiments. *, P < 0.05; **, P < 0.01; ***, P < 0.001 compared with the control in each cell type.

 
Decreased cyclin A2 and increased cyclin G1 mRNA levels were associated with Leydig cell differentiation. Because it is possible that hormonal factors regulate Leydig cell proliferative capacity by modulating cyclin levels, steady-state mRNA levels of cyclin A2 and cyclin G1 were measured in PLC, ILC, and ALC after treatment. As shown in Fig. 5Go, LH and IGF-I increased cyclin A2 mRNA levels in PLC (4.58 ± 0.66 and 4.86 ± 0.50 vs. 2.76 ± 0.21 control PLC, mean ± SE) and ILC (2.83 ± 0.32 and 2.91 ± 0.20 vs. 1.79 ± 0.14 in control ILC). Estradiol decreased cyclin A2 mRNA level in PLC (2.18 ± 0.28 vs. 2.76 ± 0.21 control PLC). Cyclin A2 mRNA levels in ALC were unaffected by any of the hormonal treatments. Treatment with LH decreased levels of cyclin G1 mRNA in PLC (0.06 ± 0.01 vs. 0.12 ± 0.02 in control) and ILC (0.31 ± 0.04 vs. 0.47 ± 0.07 in control). IGF-I decreased cyclin G1 mRNA in PLC (0.06 ± 0.01). Estradiol increased the levels of cyclin G1 mRNA in PLC (0.24 ± 0.08) and ILC (0.96 ± 0.20). MENT increased cyclin G1 mRNA in PLC (0.22 ± 0.04). Cyclin G1 mRNA levels in ALC were unaffected by any of the hormonal treatments (Fig. 6Go). These results indicated that stage-specific sets of hormonal factors regulate Leydig cell proliferative capacity during postnatal development.



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  		Figure 5. The effects of hormonal treatment on cyclin A2 mRNA levels in Leydig cells. Total cellular RNA was extracted from PLC, ILC, and ALC after 24 h culture in the absence (control) or presence of the following hormones: LH (1 ng/ml), IGF-I (70 ng/ml), MENT (50 nM), and E2 (50 nM). Steady-state levels of cyclin A2 mRNA were measured by RT-PCR after coamplification with RPS16, an internal control. Values represent the means ± SE for three separate experiments. *, P < 0.05; **, P < 0.01; ***, P < 0.001 compared with the control in each cell type.

 


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  		Figure 6. The effects of hormonal treatment on cyclin G1 mRNA levels in Leydig cells. Total cellular RNA was extracted from PLC, ILC, and ALC after 24 h culture in the absence (control) or presence of the following hormones: LH (1 ng/ml), IGF-I (70 ng/ml), MENT (50 nM), and E2 (50 nM). Steady-state levels of cyclin A2 mRNA were measured by RT-PCR after coamplification with RPS16, an internal control. Values represent the means ± SE for three separate experiments. *, P < 0.05; **, P < 0.01; ***, P < 0.001 compared with the control in each cell type.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The present study demonstrated that decreased cyclin A2 and increased cyclin G1 coincided with the loss of proliferative capacity in Leydig cells. Leydig cell progenitors had the highest proliferative capacity, and this was probably necessary for production of the pool of precursor cells from which adult Leydig cells are derived (reviewed in 1 . When Leydig cell progenitors differentiate into immature Leydig cells and subsequently into adult Leydig cells, the dramatic decline in their proliferative capacity is associated with the onset of Leydig cell differentiated function (reviewed in 1 .

Decreased cyclin A2 and increased cyclin G1 are postulated to have roles in the cessation of proliferative activity and induction of Leydig cell differentiation. The sharp decline of cyclin A2 mRNA and protein in immature and adult Leydig cells probably contributes to their lowered proliferative capacity. Cyclin A2 is thought to mediate the transition between G1 and S phase and the induction of DNA synthesis (16, 17, 18, 41). In contrast to cyclin A2, cyclin G1 increased in immature and adult Leydig cells. It has been established that the gene for cyclin G1 is activated transcriptionally by the p53 tumor suppressor protein (22, 23, 24, 25, 26). P53 is involved in cell cycle arrest, apoptosis, genomic instability, immortalization, differentiation, and stress response (reviewed in 42 . Increased cyclin G1 expression probably mediates loss of proliferative capacity, onset of differentiation, and ability to undergo apoptosis in Leydig cells.

The list of factors that modulate Leydig cell proliferation is growing, but it has not been established which of these is the primary stimulus for pubertal increases in Leydig cell numbers. However, LH is important for maintenance of high rates of Leydig cell proliferation. Administration of human CG to prepubertal rats increases the number of Leydig cells during adulthood (43) and, conversely, inactivating mutations in the LH receptor gene cause Leydig cell hypoplasia (44). In sexually mature rats, treatment with superphysiological doses of human chorionic gonadotropin for 5 weeks produced a 3-fold increase in Leydig cell number (3). These increases in adult Leydig cell numbers are dependent on proliferation and subsequent differentiation of Leydig cell precursors (45). In the present study, LH increased proliferative capacity in Leydig cell progenitors and immature Leydig cells after 1 day in vitro. The presence of LH receptors in Leydig cell progenitors and immature Leydig cells (46) indicates that LH is acting directly. The lack of a hormonal response in adult Leydig cells probably results from their terminally differentiated status, which is incompatible with cell renewal. There are reports of Leydig cell mitosis during adulthood in the guinea pig (47), mouse (48), monkeys (49), and human (50). However, adult Leydig cells in the rat fail to incorporate [3H]thymidine administered in vivo (49), nor has LH been found to stimulate their mitosis (40). This suggests that LH-induced increases in DNA synthesis during adulthood are occurring in immature Leydig cells which coexist with adult Leydig cells.

The concentration of IGF-I peaks in the interstitial fluid during puberty (6). Treatment with IGF-I in vitro is known to stimulate DNA synthesis in Leydig cell precursors (5, 51). Moreover, the number of Leydig cells in transgenic mice with a targeted deletion of the IGF-I gene was reduced to one third that of wild-type controls (52). The present study demonstrated that IGF-I stimulated proliferative capacity in Leydig cell progenitors and immature Leydig cells. A direct action of IGF-I was supported by the presence of IGF-I receptor in Leydig cell progenitors (6). Although IGF-I stimulated the proliferative capacity of Leydig cell progenitors and immature Leydig cells, it did not affect adult Leydig cells, providing further evidence of their terminally differentiated status of the adult stage.

Androgen stimulated thymidine incorporation by immature Leydig cells. The highest levels of androgen receptor mRNA and protein were observed at this stage (41, 53). Although androgen had no effect on proliferative capacity in Leydig cell progenitors, the stimulation of proliferative capacity in immature Leydig cells may explain the decreased numbers of Leydig cells in mice with testicular feminization (7, 8). Because testicular feminization in the mouse is caused by mutation that inactivates the androgen receptor (54), a role for androgen in maintaining Leydig cell numbers has been proposed. Although androgen stimulated proliferative capacity of immature Leydig cells, high levels of androgen may indirectly inhibit Leydig cell proliferation via negative suppression of LH secretion (reviewed in 55 .

Of the hormones examined in the present study, estradiol alone inhibited proliferative capacity in Leydig cell progenitors. This could explain the finding that administration of estrogen to prepubertal rats dramatically decreases adult Leydig cell numbers (10, 11). A direct action of estradiol during puberty is supported by the presence of estrogen receptor mRNA in Leydig cell progenitors at steady-state levels that are 20 times higher than adult Leydig cells (56). During the postnatal development, estradiol is secreted by Sertoli cells between days 5 and 20 (57, 58). Testicular estradiol levels decline to a nadir on day 21, then rise to a second peak during adulthood due to production by adult Leydig cells (59). Low estradiol during days 14 to 21 might allow for rapid proliferation of Leydig cell progenitors. Increased production of estradiol by immature Leydig cells and adult Leydig cells would then inhibit the further proliferation of Leydig cell progenitors, controlling the pool of Leydig cell precursors and thereby limiting Leydig cell population growth (reviewed in 60 .

Hormonal regulation of proliferative capacity in Leydig cells was hypothesized to involve cyclins. Although there are no reports with respect to LH, cyclin levels are known to be regulated by IGF-I, androgen and estrogen. IGF-I directly stimulates cyclins D1 and E (61, 62). The results herein showed that IGF-I increased cyclin A2 and decreased cyclin G1 expression. Androgen induces cyclins A, C, D1, and E, when administered exogenously, causing proliferation of prostatic glandular epithelial cells (63). Although estrogen inhibited the proliferative capacity of Leydig cells, it is an inducer of cell proliferation in some female cancers such as ovarian and breast cancers (64, 65). In the case of female cancers, estrogen is thought to act through cyclin D1 (64). The present study showed that estradiol decreased proliferative capacity in Leydig cell progenitors, and this was associated with decreased cyclin A2 and increased cyclin G1. Estradiol also increased cyclin G1 mRNA levels in immature Leydig cells, but it did not affect proliferative capacity of immature Leydig cells at this stage. This indicated that the estrogen-induced changes in cyclin G might control processes other than immature Leydig cell proliferation.

In conclusion, Leydig cells lose their proliferative capacity during postnatal differentiation. A decline in cyclin A2 and increase in cyclin G1 were associated with this loss, and hormonal factors regulated Leydig cell proliferation in a stage specific manner.


   Acknowledgments
 
We are grateful to Ms. Chantal Manon Sottas for technical assistance. We also thank Drs. Paul S. Cooke and Mary M. Lee for comments on the manuscript.


   Footnotes
 
1 Supported in part by the CONRAD program of the U.S. Agency for International Development (R.-S.G.) and NIH Grant HD-32588 (M.P.H.). Back

Received March 28, 1997.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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