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Adverse effects of di-(2-ethylhexyl) phthalate on Leydig cell regeneration in the adult rat testis
Toxicology Letters 215 (2012) 84–91
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Adverse effects of di-(2-ethylhexyl) phthalate on Leydig cell regeneration in the adult rat testis Xing-Wang Li a,1 , Yong Liang b,1 , Ying Su a,1 , Haiyun Deng a , Xiao-Heng Li a , Jingjing Guo a , Qing-Quan Lian a,∗ , Ren-Shan Ge a,c,∗ a
Institute of Reproductive Biomedicine and the 2nd Affiliated Hospital, Wenzhou Medical College, Wenzhou, Zhejiang 325027, PR China Taizhou Municipal Hospital, Medical School of Taizhou University, Jiaojiang, Zhejiang 318000, PR China c Population Council & Rockefeller University, 1230 York Avenue, New York, NY 10065, USA b
h i g h l i g h t s DEHP affects the regeneration of Leydig cells. DEHP increases the proliferation of Leydig cell precursors. DEHP increases Leydig cell number during the regeneration.
a r t i c l e
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Article history: Received 7 January 2012 Received in revised form 1 October 2012 Accepted 1 October 2012 Available online 11 October 2012 Keywords: DEHP Progenitor Leydig cells Cell proliferation Leydig cell regeneration
a b s t r a c t The objective of the present study is to determine whether di-(2-ethylhexyl) phthalate (DEHP) exposure at adulthood affects regeneration of rat Leydig cells. 90-day-old Long-Evans rats received intraperitoneal injection of 75 mg/kg ethane dimethanesulfonate (EDS) to eliminate mature Leydig cells, and then were randomly divided into 3 groups, in which rats were gavaged with the corn oil (control) or 10 or 750 mg/kg DEHP daily for 35 days. Serum testosterone and luteinizing hormone levels were assessed by RIA, Leydig cell numbers and proliferation rate were evaluated, and the mRNA levels of Leydig cell specific genes were measured by qPCR. Both 10 and 750 mg/kg DEHP treatments increased Leydig cell numbers on day 14, 21 and 35 post-EDS, due to significant increase of the number of Leydig cell precursors from day 14 to 21 post-EDS. However, serum testosterone levels were halved in 10 and 750 mg/kg DEHP groups compared to control on day 35 post-EDS despite the increased Leydig cell numbers. Quantitative PCR showed that Leydig cell specific genes including Lhcgr, Cyp11a1, Hsd3b1, and Insl3 were significantly down-regulated in 750 mg/kg DEHP-treated testes on post-EDS day 21 and beyond. The present study suggests that DEHP increases Leydig cell proliferation but inhibits differentiation during the regeneration of Leydig cells. © 2012 Elsevier Ireland Ltd. All rights reserved.
1. Introduction Phthalates are a class of dialkyl or alkyl aryl esters of 1,2benzenedicarboxylic acid. Phthalates are used as plasticizers in polyvinyl chloride plastics that make consumer products such as flexible plastic and toys, curtains, wallpaper, food packaging, and plastic wrap and medical devices. Phthalates are also used in cosmetics and personal care products, including perfume, soap, shampoo, hair spray, nail polish, and skin moisturizers. One of most abundantly used phthalates is di-(2-ethylhexyl) phthalate (DEHP).
∗ Corresponding authors at: The 2nd Affiliated Hospital, Wenzhou Medical College, Wenzhou, Zhejiang 325027, PR China. Tel.: +86 577 88879079. E-mail addresses: [email protected] (Q.-Q. Lian), r [email protected] (R.-S. Ge). 1 These authors equally contributed to the work. 0378-4274/$ – see front matter © 2012 Elsevier Ireland Ltd. All rights reserved. http://dx.doi.org/10.1016/j.toxlet.2012.10.001
Because more than 95% human urinary samples have been detected to contain its metabolites (Meeker et al., 2007; Zhang et al., 2009), DEHP is believed to be ubiquitously present in human being. The United States Environmental Protection Agency (EPA) has classified DEHP as a chemical that may be potentially harmful to human health. Several studies indicate an association of phthalates with some potentially adverse endpoints, including interference with the timing of parturition, shortening anogenital distance of boys and lowering birth weight of neonates (Adibi et al., 2009; Swan, 2008; Zhang et al., 2006, 2009). In laboratory animals, DEHP has been reported to cause a dose-dependent increase in the incidence of Leydig cell tumors in adult Sprague Dawley rats when they were exposed to 0–300 mg/kg DEHP per day for 159 weeks (Voss et al., 2005). DEHP also induced a shortening of anogenital distance and lower testicular testosterone level of male pups in the postnatal day 2–3 when dams were exposed to 750 mg/kg DEHP from gestational day 14 to postnatal day 3 (Gray et al., 2000). Our previous study
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demonstrated that DEHP exposure during pubertal periods caused Leydig cell hyperplasia in rats (Akingbemi et al., 2001, 2004). Due to the proliferative capacity of Leydig cell precursors during puberty, increased mitosis of these precursor cells may be the cause of Leydig cell hyperplasia in rats (Akingbemi et al., 2001, 2004). However, there is still unclear whether DEHP exposure affects the Leydig cell functions at adulthood. In the adult testis, a small pool of stem cells regenerates to compensate the loss of mature Leydig cells. However, it is clear from studies of adult rats that the resident stem cells are able to support the restoration of adult populations of Leydig cells to testes from which the originally present cells have been experimentally eliminated by the alkylating agent ethane dimethane sulfonate (EDS) (Teerds, 1996). The loss of Leydig cells triggers a rapid regeneration of new adult population of Leydig cells within 4 weeks in the rat testis due to high circulatory level of luteinizing hormone (LH) (Kerr et al., 1985). Four days after the treatment of EDS, all Leydig cells are eliminated, and 14 days post-EDS newly formed progenitor Leydig cells appear and they gradually differentiate into more mature Leydig cells in this lineage from 21 to 35 days post-EDS (Teerds, 1996). In the present study, we reported the effects of DEHP exposures on the regeneration process of new population of Leydig cells at adulthood in EDS-treated model. 2. Materials and methods 2.1. Animal treatment The EDS was a gift from Dr. L. Earl Gray (U.S. Environmental Protection Agency, Research Triangle Park, NC). EDS was dissolved in a mixture of dimethyl sulfoxide (DMSO) and water (DMSO:H2 O, 1:4, v/v) for intraperitoneal injection. Twenty-four 90-day-old male Long-Evans rats (Charles River, Raleigh, NC) received a vehicle control (DMSO:H2 O, 1:4, v/v), and six rats per time-point were killed at days 4, 14, 21 and 35 post-EDS as the control. Seventy-two 90-day-old male Long-Evans rats (Charles River, Raleigh, NC) received a single intraperitoneal injection of 75 mg/kg EDS to eliminate Leydig cells, and then randomly divided into three groups with 24 rats that were gavaged with vehicle (corn oil 1.0 ml/250 g rat), 24 rats that were gavaged with 10 mg/kg/day DEHP (Sigma, CAS 204-211-0, purity of 99%) and 24 rats that were gavaged with 750 mg/kg/day DEHP. DEHP was gavaged 1 h after EDS for 35 days. DEHP was suspended in corn oil and gavaged to the rat by 0.5 ml/250 g rat. Dosages (0, 10, or 750 mg/kg per day DEHP) were selected based on our previously determined lowest-observed-effect level (10 mg/kg) of DEHP that increases Leydig cell number (Akingbemi et al., 2001; Ge et al., 2007), and lowest-observed-effect level (750 mg/kg) of DEHP that inhibits serum testosterone level in the pubertal rats (Foster et al., 2001; Ge et al., 2007). Eight rats per group were euthanized by CO2 on days 14, 21 and 35 post-EDS. Two hours before they were killed, all animals received an intraperitoneal injection of BrdU (40 g/g body weight, Cat. No. 280879 Roche Diagnostics) to label dividing cells. Blood samples were collected, placed in a gel tube and centrifuged at 1500 × g for 10 min to collect serum. The serum was stored at −20 ◦ C until the radioimmunoassay for T and LH. One testis from each animal was removed and frozen in liquid N2 for subsequent study of mRNA and steroidogenic enzyme levels, and the contralateral testis from each animal was punched holes using a needle and fixed by Bouin’s solution. All animal procedures were performed in accordance with the policies of The Rockefeller University’s Animal Care and Use Committee. 2.2. Serum testosterone and LH analysis Serum testosterone concentrations were measured with a tritium-based radioimmunoassay as previously described (Ge and Hardy, 1998). Interassay variation of the testosterone assay was between 7 and 8%. Serum LH concentrations were measured using the method described by Chandrashekar and Bartke (1988). LH standards and antibody were obtained through the National Hormone and Pituitary Program. Radioactive 125 I-rat LH was obtained through Covance Laboratories (Vienna, VA), and IgG antiserum was obtained from ICN Pharmaceuticals (Costa Mesa, CA). The sensitivity and inter- and intra-assay coefficients of variation for the LH assay were, respectively, 0.08 ng per ml and 5% and 6%.
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(Payne et al., 1980). After 3HSD reaction, nitro-blue tetrazoliumas is reduced and deposited as a diformazan at the site of Leydig cell cytosol. 4 ,6-Diamidino2-phenylindole (DAPI) was used as nuclear counterstaining of a cell. Testis sections were captured by using a Nikon Eclipse E800 microscope (Nikon) and a SPOT RT digital camera (model 2.3.0.; Diagnostic Instruments) interfaced to a computer. Cell numbers were estimated using image analysis software (Image-Pro Plus, Media Cybernetics, Rockville, MD). Leydig cell numbers were enumerated according to the fractionator technique (Akingbemi et al., 2004). Testes were cut and sections were collected every 15 section cuts for further analysis. About 10 sections were sampled randomly from each testis. The total number of Leydig cells per 100 seminiferous tubules was calculated since the tubule number was not affected by EDS. 2.4. BrdU incorporation Four testes per group per time-point were randomly selected. The testes were then removed, punched three holes per testis with needle to allow fixative to flow in and stored in the Bouin solution fixative for two days. After dehydration in ethanol and xylene, the testes were embedded in paraffin for immunocytochemical analysis. Leydig cells were identified through immunopositive staining for the marker enzyme 3HSD. In brief, 6-mm-thick transverse sections were prepared and mounted on glass slides (Cat. No. 12-550-15; Fisher Scientific Company, Hampton, NH). Avidin-biotin immunostaining was performed using a kit (Cat. Nos. PK-2200 for BrdU and PK-6101 for 3HSD; Vector Laboratories Inc., Burlingame, CA) according to the manufacturer’s instructions. Antigen retrieval was carried out by microwave irradiation for 10 min in 10 mM (pH 6.0) citrate buffer, and endogenous peroxidase was blocked with 0.5% H2 O2 in methanol for 30 min. The sections were then incubated with a monoclonal anti-BrdU antibody (RPN 202; Amersham Biosciences, Little Chalfont, UK) for 30 min at room temperature. The antibody bound to the nuclei was visualized with diaminobenzidine (Cat. No. sk-4100; Vector Laboratories Inc.) and the labeled nuclei were stained black by adding a nickel solution to the chromogen. After washing, the sections were double-labeled by incubation with a 3HSD polyclonal antibody diluted 1:3000 (provided by Dr. Van Luu-The, Laval University, Quebec, Canada) for 1 h at room temperature. The antibody–antigen complexes were visualized with diaminobenzidine alone, resulting in brown cytoplasmic staining in positively labeled Leydig cells. The sections were counterstained with Mayer hematoxylin, dehydrated in graded concentrations of alcohol, and cover-slipped with resin (Permount, SP15-100; Fisher Scientific). In control experiments, sections were incubated with nonimmune rabbit IgG (3HSD) or mouse IgG (BrdU) using the same working dilution as the primary antibody. 16 mosaic sections for each of three nonadjacent sections per testis were captured using a Nikon Eclipse E800 microscope (Nikon, Inc., Melville, NY) equipped with a 60 objective lens and a SPOT RT digital camera (model 2.3.0., Diagnostic Instruments, Inc., Michigan City, IN) interfaced to a computer. 200–300 Leydig cells were enumerated. Labeling index (LI) was calculated as BrdU positive cells per 100 Leydig cells. 2.5. Quantitative PCR Six testes per group per time-point were randomly selected. Total RNA was extracted from rat testes in Trizol according to the manufacturer’s instructions (Invitrogen, Carlsbad, CA). First-strand synthesis and real-time PCR (qPCR) were performed as previously described (Ge et al., 2005). Ribosomal protein S16 (Rps16) mRNA levels were assayed in all samples as an internal control. The expression levels of nine Leydig cell specific genes analyzed and the primers for the following genes were described in our previous study (Lin et al., 2008). These genes include receptor gene: luteinizing hormone receptor (Lhcgr); steroidogenic transporters and enzymes: scavenger receptor class B member 1 (Scarb1), steroidogenic acute regulatory protein (Star), cholesterol side chain cleavage enzyme (CYP11A1, encoded by Cyp11a1), 3HSD1 (Hsd3b1), 17␣-hydroxylase/20lyase (CYP17A1, encoded by Cyp17a1) and 17-hydroxysteroid dehydrogenase 3 (17HSD3, encoded by Hsd17b3); biomarker for Leydig cells at the more advanced stage: 11-hydroxysteroid dehydrogenase 1 (Hsd11b1) (Phillips et al., 1989); and Leydig cell differentiation status marker: insulin-like growth factor 3 (Insl3). 2.6. Homogenization and protein content assay The testes (randomly selected 6 samples per group per time-point) were homogenized in 1 ml ice-cold 0.1 M PBS (pH 7.2) containing 0.25 M sucrose. Supernatants were collected by centrifugation at 700 × g for 30 min. Supernatants were used to measure the protein level of CYP11A1, and enzyme activities of CYP17A1, 3HSD1 and 17HSD3. The protein concentrations were determined using a kit (No. 5000006, Bio-Rad Laboratories, Inc., Hercules, CA) with BSA as a standard. 2.7. Western blot analysis of CYP11A1
2.3. Determination of Leydig cell number Four testes per group per time-point were randomly selected. Frozen testis was sectioned in a cryostat, and 8 m testis sections were cut. Leydig cells were identified by the histochemical staining of Leydig cell specific marker 3-hydroxysteroid dehydrogenase (3HSD) activity with 0.4 mM etiocholanolone and 1 mg nitro-blue tetrazoliumas and NAD+ as cofactor according to the previously described method
Protein (50 g) was boiled in equal volumes of sample loading buffer, a Tris–Cl buffer (pH 6.8) containing 20% glycerol, 5% SDS, 3.1% dithiothreitol, and 0.001% bromophenol blue. Protein samples were electrophoresed on 10% polyacrylamide gels containing SDS. Proteins were electrophoretically transferred onto nitrocellulose membranes, and after 30 min exposure to 10% nonfat milk to block nonspecific binding, the membrane was incubated with a CYP11A1 antibody (catalogue no.
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RDIp450sccabr, RDI Research Diagnostics, Inc., Flanders, NJ; 1:1000). The membranes were then washed and incubated with a 1:5000 dilution of second antibody that was conjugated to horseradish peroxidase (anti-rabbit Ig, HRP-linked whole antibody produced in donkey; Amersham Biosciences Corp., Piscataway, NJ). The washing step was repeated, and immunoreactive bands were visualized by chemiluminescence using a kit (ECL, Amersham, Arlington Heights, IL). Then the antibody in the membrane was stripped away and probed again by an antibody against -actin (ACTB, Sigma; no. A2228; 1:1000). The second anti-mouse Ig, HRP-linked whole antibody produced in sheep (Amersham Biosciences, Piscataway, NJ) was used and the ACTB band was visualized by chemiluminescence. Protein levels were measured by densitometry of the films and normalized to ACTB. 2.8. Enzyme assay Testosterone biosynthetic enzymes activities of CYP17A1, 3HSD1 and 17HSD3 were determined by thin layer chromatography (TLC) and radiometry as previously described (Ye et al., 2011). The reaction mixtures (total volume of 250 l) containing 25–160 g protein, 0.2 mM cofactors (NAD+ for 3HSD1, NADPH for CYP17A1, and 17HSD3), and 1000 nM steroid substrates (radiolabeled plus cold substrates) were incubated in shaking water bath at 34 C for 1–3 h. The substrates were follows: pregnenolone (for 3HSD1), progesterone (for CYP17A1), and androstenedione (for 17HSD3). The preliminary experiment was conducted to determine the linear reaction curve using different concentrations of proteins at different time periods. The steroids were extracted from reaction mixture with 1 ml of ice-cold ether, and the organic layer was evaporated under nitrogen gas. The extract of steroids was suspended in 70 l ether and then spotted on thin layer plates (Baker-flex, Phillipsburg, NJ). The steroids were separated chromatographically in chloroform:methanol (97:3, v/v) for 3HSD1, and 17HSD3 as well as chloroform:ether (7:1, v/v) for CYP17A1. The radioactivity was measured with a scanning radiometer (System 200/AC3000, Bioscan, Inc., Washington DC). The conversion of steroid to product was calculated as a percentage of the total radioactivity found in the product. All assays were repeated in triplicate. 2.9. Statistical analysis Values are expressed as mean ± SEM, and data were analyzed using one-way ANOVA with Tukey’s comparison of all groups at each time point post-EDS using GraphPad Prism (version 5, GraphPad Software Inc., San Diego, CA). Significant difference was regarded at P < 0.05.
3. Results 3.1. Gestational exposure 3.1.1. General toxicological parameters and serum testosterone and LH levels Gavage of rats with 10 and 750 mg/kg DEHP for 35 days did not affect body weights (data not shown). After DEHP treatment, all rats were survived and had normal clinical signs, confirming the results of our previous study. Serum testosterone level was used as an indicator of the regeneration process. The baseline of testosterone level in the adult Long Evens male rats without EDS was about 5 ng/ml (Akingbemi et al., 2001). Serum testosterone levels were significantly reduced by 10 and 750 mg/kg DEHP on day 35 post-EDS (Fig. 1A). Serum LH level was significantly higher in the 10 mg/kg DEHP group on day 21 post-EDS. 3.1.2. Effects of DEHP on Leydig cell number and proliferation during regeneration Leydig cells can be identified by 3HSD staining (blue diformazan deposit). 3HSD staining reflects the Leydig cells but the status of Leydig cell differentiation. As shown in Fig. 2A, there were many 3HSDpos cells in the interstitium of 90-day-old rat testis. Four days post-EDS, all 3HSDpos cells were eliminated (Fig. 2B). 14 days post-EDS, there was one or 2 newly formed 3HSDpos cells in the control testis (Fig. 2C), while there were increased numbers of 3HSDpos cells in the DEHP-treated testes (Fig. 2D illustrates the 3HSDpos cells in the 10 mg/kg DEHP-treated testis). Enumeration of 3HSDpos cells showed that there were significantly and dose-dependently increased numbers of 3HSDpos cells in both DEHP-treated groups on day 14 (Fig. 3A), 21 (Fig. 3B) and 35 (Fig. 3C) post-EDS. The numbers of 3HSDpos cells in 10 and 750 mg/kg DEHP
Fig. 1. Serum testosterone and LH levels after DEHP exposure during the course of regeneration. Rats were i.p. treated 75 mg/kg EDS and 1 h later they were gavaged with 0 or 10 or 750 mg DEHP. Panel A: testosterone, and Panel B: LH. Bar , and represents 0 (control), 10 and 750 mg/kg DEHP, respectively. Mean ± SEM, n = 6, *significant change compared to control at each time point.
testes compared to control were increased by 3.5 and 9 fold on day 14. However, the number of 3HSDpos cells in both DEHP-treated groups was only about 1.36 fold of control on day 35 post-EDS. This indicates that DEHP treatment promotes the proliferation of 3HSDpos cells during earlier stage of regeneration. We further investigated the proliferation of Leydig cells using BrdU incorporation. As shown in Fig. 4, 10 and 750 mg/kg DEHP significantly increased the labeling index of Leydig precursor cells during regeneration on days 14 and 21 post-EDS. However, labeling index of Leydig cells never reached significance of both DEHPtreated groups when compared to the vehicle control on day 35 post-EDS (Fig. 4C). This indicates that the increased numbers of Leydig cells are mainly from the Leydig precursor cells at the earlier stage of regeneration. 3.1.3. Effects of DEHP on Leydig cell differentiation during regeneration Although there was the increased number of Leydig cells during the regeneration by DEHP, serum testosterone levels in both DEHP-treated groups were only half of that of control on day 35 post-EDS, indicating that there is defect of Leydig cell
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Fig. 2. Histochemical staining of 3HSD1 of testis. 3HSD1 was stained blue, and nucleus was fluorescently stained white by DAPI. Rats were i.p. treated 75 mg/kg EDS and 1 h later they were gavaged with 0 or 10 or 750 mg DEHP. Panel A: normal 94-day-old testis; Panel B: 14-day-post-EDS control testis; Panel C: 10 mg/kg DEHP-treated testis on day 14 post-EDS; and Panel D: 750 mg/kg DEHP-treated testis on day 14 post-EDS. Black arrows point to 3HSD1 positive cells. Bar = 50 m. (For interpretation of the references to color in Figure legend, the reader is referred to the web version of the article.)
differentiation in DEHP-treated testis. The expression levels of several Leydig cell-specific genes as biomarkers of differentiated status of Leydig cells during regeneration were shown in Fig. 5. We found that although Leydig cell numbers were increased by 3.5 and 9 fold in 10 and 750 mg/kg DEHP-treated testes on day 14 post-EDS, the expression levels of all selected Leydig cell specific genes were not altered. On day 21 post-EDS, only Star reached significant increase in 10 mg/kg DEHP group, while Insl3, a gene that reflects the differentiation status of Leydig cells and is not regulated by LH,
Leydig cell #/100 tubules
A. 14 days 600
was significantly decreased in both 10 and 750 mg/kg DEHP-treated testes. At that time point, the expression levels of steroidogenic enzyme genes Cyp11a1 and Hsd3b1 were significantly decreased in 750 mg/kg DEHP-treated testes. On day 35 post-EDS, the expression levels of Lhcgr, Cyp11a1, Hsd3b1 and Hsd11b1 were down-regulated in both 10 and 750 mg/kg DEHP-treated testes. We determined the protein levels of CYP11A1 and enzyme activities of CYP17A1, 3HSD1 and 17HSD3 on day 35 post-EDS, and found that CYP11A1 level and 3HSD1 activity were significantly
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Fig. 3. Leydig cell number. Leydig cell numbers (#) were calculated as cells per 100 tubules. Mean ± SEM (n = 4). Rats were i.p. treated 75 mg/kg EDS and 1 hr later they were gavaged with 0 or 10 or 750 mg DEHP. Panel A: 14 days; Panel B: 21 days; and Panel C: 35 days post-EDS. *, **Significant differences compared to control at P < 0.05 and 0.01 at each time point post-EDS, respectively.
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Labeling index (100%)
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Fig. 4. Labeling index of Leydig cells. Labeling index was calculated as BrdU positive cells per 100 Leydig cells. Rats were i.p. treated 75 mg/kg EDS and 1 h later they were gavaged with 0 or 10 or 750 mg DEHP. Mean ± SEM (n = 4). Panel A: 14 days; Panel B: 21 days; and Panel C: 35 days post-EDS. Labeling index was calculated by the percentage of BrDU-positive cells per 200 interstitial cells. *, **Significant differences compared to control at P < 0.05 and 0.01 at each time point post-EDS, respectively.
reduced in both doses of DEHP while CYP17A1 and 17HSD3 were without changes (Fig. 6), confirming their mRNA level changes. 4. Discussion The EDS-treated rat Leydig cell regeneration model is very unique for observing the proliferation and differentiation of Leydig cells in the adult testis. Leydig cells in the adult testis are completely eliminated after a single intraperitoneal injection of EDS (Teerds, 1996). The loss of Leydig cells leads to the elevation of circulating LH, which together with local growth factors or cytokines promotes the proliferation of Leydig precursor cells and thereafter differentiation. The present study demonstrated that DEHP exposure to adult rats led to the increased Leydig cell number due to the increased the proliferative capacity of these newly formed Leydig precursor cells during the earlier regeneration process (from days 14 to 21 postEDS). Stereological enumeration of Leydig (3HSDpos ) cell numbers showed that the earlier exposure to DEHP have caused 3.5 and 9 fold on day 14, 1.5 and 1.8 fold on day 21, as well as 1.36 and 1.36 fold increase in Leydig cell numbers on day 35 post-EDS (Fig. 3). Apparently, the increased number of Leydig cells came from the proliferation of Leydig cell precursors from day 14 to 21 post-EDS (Fig. 4). The lack of significance for BrDU labeling at day 35 postEDS in not merely be a representation that spatial capacity of the interstitial compartment was reached The regeneration of Leydig cells showed a very similar pattern of pubertal Leydig cell development with higher proliferative rate in progenitor Leydig cells and lower proliferative rate later on (Ge and Hardy, 1997). The testis weight post-EDS treatment cannot reflect the Leydig cells because the testis weight was significantly reduced after EDS due to blocked spermatogenesis after loss of testosterone. Quite interestingly, high dose of DEHP exposure to adult rats inhibited the differentiation of Leydig cells, as judged by the decreased expression levels of Cyp11a1, Hsd3b1, Lhcgr, and Insl3, starting on day 21 post-EDS. Interestingly, Insl3, a gene that reflects the differentiation status of Leydig cells or Leydig cell numbers and is not regulated by LH, was also significantly decreased in low dose (10 mg/kg) and high dose (750 mg/kg) DEHP-treated testes. Since the Leydig cell numbers were actually increased in both doses of DEHP, the decreased expression level of Insl3 may reflect the delayed status of Leydig cell differentiation at 21 days post-EDS. Even at 35-day post EDS, the Leydig cell numbers were doubled in both doses of DEHP, the reduction of Insl3 expression levels in these groups indicates that the delayed differentiation of Leydig cells even at 35 days post-EDS. The serum level of testosterone
usually reflects the differentiated status of Leydig cells or the Leydig cell numbers in the testis. Apparently, this level reached significant decrease on day 35 post-EDS (Fig. 1A). There were no significant changes of serum testosterone levels from days 14 to 21 post-EDS, this might be due to significant higher ratio of number of Leydig precursor cells on days 14 (3.5–9 fold) and 21 (1.5 to 1.8 fold). The expression levels of Lhcgr, Hsd3b1 and Hsd11b1 in the DEHP-treated testes were decreased, even the Leydig cell numbers were increased, indicating that these Leydig cells may be less differentiated and produce fewer amount of testosterone. Other researchers also reported that the decreases in serum testosterone at high doses of DEHP when administered from weaning through adulthood without EDS administration (Noriega et al., 2009). The expression levels of Leydig cell specific genes, including Cyp11a1, Hsd3b1 were actually decreased though such higher ratio of Leydig cell numbers in high-dose DEHP-treated groups vs. the control group. The lowest observed adverse effect level (LOAEL) for increasing Leydig cell number and decreasing the differentiation of Leydig cells was 10 mg/kg/day. This level is within the suggested environmental exposure levels of the human population. Previously, our laboratory and others have demonstrated that low dose of DEHP can increase Leydig cell numbers during puberty by increasing the proliferation of Leydig precursor cells (Akingbemi et al., 2001, 2004). It is true that during puberty Leydig cell precursor cells (progenitor and immature Leydig cells) are proliferative (Ge and Hardy, 1998). Specifically, progenitor Leydig cells have higher proliferative capacity. However, at adulthood, cell replication slowed abruptly after puberty. The estimated turnover time for adult Leydig cells exceeds the two-year lifespan of the average rat (Kerr et al., 1987). In the present study, we also observed a significant increase of proliferation of Leydig cells during the earlier regeneration process in EDS-treated model, thus contributing to a significantly increased Leydig cell numbers. A recent study also showed that high dose of dibutyl phthalate, another widely used phthalate, significantly increased Leydig cell numbers during regeneration after EDS (Heng et al., 2011). This indicates that dibutyl phthalate and DEHP have a similar mechanism. However, the mechanism of stimulating Leydig cell proliferation in the regeneration model may be different from that caused by DEHP during pubertal exposure. Our previous study observed a significant increase LH levels by DEHP during pubertal exposure (Akingbemi et al., 2001, 2004). LH level is controlled by the circulating androgen. The normal adult Long Evens rats had serum LH levels around 0.5 ng/ml (Akingbemi et al., 2001). Loss of Leydig cells after EDS treatment significantly increased circulating LH level, and at 14 days post-EDS serum LH levels were still around 10 ng/ml
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Fig. 5. Steady-state mRNA levels of Leydig cell specific genes after DEHP treatment by quantitative PCR. Rats were i.p. treated 75 mg/kg EDS and 1 h later they were gavaged with 0 or 10 or 750 mg DEHP. Mean ± SEM (n = 5). (a)–(c) indicate significant differences of each DEHP-treated group compared to control (0 mg/kg DEHP) at each time point post-EDS at P < 0.05, 0.01, 0.001, respectively.
(Fig. 1B). In the present study, we only observed the increased LH level in the 10 mg/kg DEHP group on day 21 post-EDS, although there were significant increases of Leydig cell proliferation in both 10 and 750 mg/kg DEHP-treated groups. This difference may be complicated by the elevated LH levels in EDS-treated rats due to the negative feedback after elimination of Leydig cells. The change of LH levels may not be important factor to contribute to the proliferation of Leydig cell precursors, since the initial proliferation and
differentiation of Leydig cell precursors can occur in the absence of LH in hypogonadal rats (Teerds et al., 1989). In conclusion, the present study shows that DEHP exposure at adulthood increased Leydig cell numbers but inhibited the differentiation process during the regeneration. Leydig cell hyperplasia has been documented in rat neonates following in utero exposure to DEHP (Lin et al., 2008; Mahood et al., 2005; Mylchreest et al., 2002; Parks et al., 2000).
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Fig. 6. Protein level and steroidogenic enzyme activity of testis after DEHP treatment. CYP11A1 was measured by Western blotting (Panel A and B). Rats were i.p. treated 75 mg/kg EDS and 1 h later they were gavaged with 0 or 10 or 750 mg DEHP. 3HSD1 was measured as described as in Section 2. Mean ± SEM (n = 5–6). **, ***Significant difference of each DEHP-treated group compared to control (0 mg/kg DEHP) at P < 0.01 and 0.001, respectively.
Conflict of interest statement None. Acknowledgements We thank Chantal M. Sottas for the technical support from the Population Council. This work was also partially funded by NSFC 30871434 (R.S.G.) and Wenzhou Science & Technology Funding Project H20090075 (X. W.L.). Partial data were presented in American Society Annual Meeting held in 2009. References Adibi, J.J., Hauser, R., Williams, P.L., Whyatt, R.M., Calafat, A.M., Nelson, H., Herrick, R., Swan, S.H., 2009. Maternal urinary metabolites of di-(2-ethylhexyl) phthalate
in relation to the timing of labor in a US multicenter pregnancy cohort study. American Journal of Epidemiology 169, 1015–1024. Akingbemi, B.T., Ge, R., Klinefelter, G.R., Zirkin, B.R., Hardy, M.P., 2004. Phthalateinduced Leydig cell hyperplasia is associated with multiple endocrine disturbances. Proceedings of the National Academy of Sciences of the United States of America 101, 775–780. Akingbemi, B.T., Youker, R.T., Sottas, C.M., Ge, R., Katz, E., Klinefelter, G.R., Zirkin, B.R., Hardy, M.P., 2001. Modulation of rat Leydig cell steroidogenic function by di(2-ethylhexyl)phthalate. Biology of Reproduction 65, 1252–1259. Chandrashekar, V., Bartke, A., 1988. Influence of endogenous prolactin on the luteinizing hormone stimulation of testicular steroidogenesis and the role of prolactin in adult male rats. Steroids 51, 559–576. Foster, P.M., Mylchreest, E., Gaido, K.W., Sar, M., 2001. Effects of phthalate esters on the developing reproductive tract of male rats. Human Reproduction Update 7, 231–235. Ge, R.S., Chen, G.R., Dong, Q., Akingbemi, B., Sottas, C.M., Santos, M., Sealfon, S.C., Bernard, D.J., Hardy, M.P., 2007. Biphasic effects of postnatal exposure to diethylhexylphthalate on the timing of puberty in male rats. Journal of Andrology 28, 513–520. Ge, R.S., Dong, Q., Niu, E.M., Sottas, C.M., Hardy, D.O., Catterall, J.F., Latif, S.A., Morris, D.J., Hardy, M.P., 2005. 11beta-Hydroxysteroid dehydrogenase 2 in rat leydig cells: its role in blunting glucocorticoid action at physiological levels of substrate. Endocrinology 146, 2657–2664. Ge, R.S., Hardy, M.P., 1997. Decreased cyclin A2 and increased cyclin G1 levels coincide with loss of proliferative capacity in rat Leydig cells during pubertal development. Endocrinology 138, 3719–3726. Ge, R.S., Hardy, M.P., 1998. Variation in the end products of androgen biosynthesis and metabolism during postnatal differentiation of rat Leydig cells. Endocrinology 139, 3787–3795. Gray L.E. Jr., Ostby, J., Furr, J., Price, M., Veeramachaneni, D.N., Parks, L., 2000. Perinatal exposure to the phthalates DEHP BBP and DINP but not DEP DMP or DOTP alters sexual differentiation of the male rat. Toxicological Sciences 58, 350– 365. Heng, K., Anand-Ivell, R., Teerds, K., Ivell, R., 2011. The endocrine disruptors dibutyl phthalate (DBP) and diethylstilbestrol (DES) influence Leydig cell regeneration following ethane dimethane sulphonate treatment of adult male rats. International Journal of Andrology 35, 353–363. Kerr, J.B., Donachie, K., Rommerts, F.F., 1985. Selective destruction and regeneration of rat Leydig cells in vivo. A new method for the study of seminiferous tubular-interstitial tissue interaction. Cell and Tissue Research 242, 145– 156. Kerr, J.B., Knell, C.M., Irby, D.C., 1987. Ultrastructure and possible function of giant crystalloids in the Sertoli cell of the juvenile and adult koala (Phascolarctos cinereus). Anatomy and Embryology 176, 213–224. Lin, H., Ge, R.S., Chen, G.R., Hu, G.X., Dong, L., Lian, Q.Q., Hardy, D.O., Sottas, C.M., Li, X.K., Hardy, M.P., 2008. Involvement of testicular growth factors in fetal Leydig cell aggregation after exposure to phthalate in utero. Proceedings of the National Academy of Sciences of the United States of America 105, 7218–7222. Mahood, I.K., Hallmark, N., McKinnell, C., Walker, M., Fisher, J.S., Sharpe, R.M., 2005. Abnormal Leydig cell aggregation in the fetal testis of rats exposed to di (nbutyl) phthalate and its possible role in testicular dysgenesis. Endocrinology 146, 613–623. Meeker, J.D., Calafat, A.M., Hauser, R., 2007. Di(2-ethylhexyl) phthalate metabolites may alter thyroid hormone levels in men. Environmental Health Perspectives 115, 1029–1034. Mylchreest, E., Sar, M., Wallace, D.G., Foster, P.M., 2002. Fetal testosterone insufficiency and abnormal proliferation of Leydig cells and gonocytes in rats exposed to di(n-butyl) phthalate. Reproductive Toxicology 16, 19–28. Noriega, N.C., Howdeshell, K.L., Furr, J., Lambright, C.R., Wilson, V.S., Gray Jr., L.E., 2009. Pubertal administration of DEHP delays puberty, suppresses testosterone production, and inhibits reproductive tract development in male Sprague-Dawley and Long-Evans rats. Toxicological Sciences 111, 163–178. Parks, L.G., Ostby, J.S., Lambright, C.R., Klinefelter, G.R., Abbott, B.D., Barlow, N.J., Gray Jr., L.E., 2000. The plasticizer diethylhexyl phthalate induces malformations by decreasing fetal testosterone synthesis during sexual differentiation in the male rat. Toxicological Sciences 58, 339–349. Payne, A.H., Downing, J.R., Wong, K.L., 1980. Luteinizing hormone receptors and testosterone synthesis in two distinct populations of Leydig cells. Endocrinology 106, 1424–1429. Phillips, D.M., Lakshmi, V., Monder, C., 1989. Corticosteroid 11 -dehydrogenase in rat testis. Endocrinology 125, 209–216. Swan, S.H., 2008. Environmental phthalate exposure in relation to reproductive outcomes and other health endpoints in humans. Environmental Research 108, 177–184. Teerds, K.J., 1996. Regeneration of Leydig cells after depletion by EDS: a model for postnatal Leydig cell renewal. In: Payne, A.H., Hardy, M.P., Russell, L.D. (Eds.), The Leydig Cell. Cache River Press, Vienna, IL, pp. 203–220. Teerds, K.J., de Rooij, D.G., Rommerts, F.F., van den Hurk, R., Wensing, C.J., 1989. Proliferation and differentiation of possible Leydig cell precursors after destruction of the existing Leydig cells with ethane dimethyl sulphonate: the role of LH/human chorionic gonadotrophin. Journal of Endocrinology 122, 689– 696.
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Toxicology Letters journal homepage: www.elsevier.com/locate/toxlet
Adverse effects of di-(2-ethylhexyl) phthalate on Leydig cell regeneration in the adult rat testis Xing-Wang Li a,1 , Yong Liang b,1 , Ying Su a,1 , Haiyun Deng a , Xiao-Heng Li a , Jingjing Guo a , Qing-Quan Lian a,∗ , Ren-Shan Ge a,c,∗ a
Institute of Reproductive Biomedicine and the 2nd Affiliated Hospital, Wenzhou Medical College, Wenzhou, Zhejiang 325027, PR China Taizhou Municipal Hospital, Medical School of Taizhou University, Jiaojiang, Zhejiang 318000, PR China c Population Council & Rockefeller University, 1230 York Avenue, New York, NY 10065, USA b
h i g h l i g h t s DEHP affects the regeneration of Leydig cells. DEHP increases the proliferation of Leydig cell precursors. DEHP increases Leydig cell number during the regeneration.
a r t i c l e
i n f o
Article history: Received 7 January 2012 Received in revised form 1 October 2012 Accepted 1 October 2012 Available online 11 October 2012 Keywords: DEHP Progenitor Leydig cells Cell proliferation Leydig cell regeneration
a b s t r a c t The objective of the present study is to determine whether di-(2-ethylhexyl) phthalate (DEHP) exposure at adulthood affects regeneration of rat Leydig cells. 90-day-old Long-Evans rats received intraperitoneal injection of 75 mg/kg ethane dimethanesulfonate (EDS) to eliminate mature Leydig cells, and then were randomly divided into 3 groups, in which rats were gavaged with the corn oil (control) or 10 or 750 mg/kg DEHP daily for 35 days. Serum testosterone and luteinizing hormone levels were assessed by RIA, Leydig cell numbers and proliferation rate were evaluated, and the mRNA levels of Leydig cell specific genes were measured by qPCR. Both 10 and 750 mg/kg DEHP treatments increased Leydig cell numbers on day 14, 21 and 35 post-EDS, due to significant increase of the number of Leydig cell precursors from day 14 to 21 post-EDS. However, serum testosterone levels were halved in 10 and 750 mg/kg DEHP groups compared to control on day 35 post-EDS despite the increased Leydig cell numbers. Quantitative PCR showed that Leydig cell specific genes including Lhcgr, Cyp11a1, Hsd3b1, and Insl3 were significantly down-regulated in 750 mg/kg DEHP-treated testes on post-EDS day 21 and beyond. The present study suggests that DEHP increases Leydig cell proliferation but inhibits differentiation during the regeneration of Leydig cells. © 2012 Elsevier Ireland Ltd. All rights reserved.
1. Introduction Phthalates are a class of dialkyl or alkyl aryl esters of 1,2benzenedicarboxylic acid. Phthalates are used as plasticizers in polyvinyl chloride plastics that make consumer products such as flexible plastic and toys, curtains, wallpaper, food packaging, and plastic wrap and medical devices. Phthalates are also used in cosmetics and personal care products, including perfume, soap, shampoo, hair spray, nail polish, and skin moisturizers. One of most abundantly used phthalates is di-(2-ethylhexyl) phthalate (DEHP).
∗ Corresponding authors at: The 2nd Affiliated Hospital, Wenzhou Medical College, Wenzhou, Zhejiang 325027, PR China. Tel.: +86 577 88879079. E-mail addresses: [email protected] (Q.-Q. Lian), r [email protected] (R.-S. Ge). 1 These authors equally contributed to the work. 0378-4274/$ – see front matter © 2012 Elsevier Ireland Ltd. All rights reserved. http://dx.doi.org/10.1016/j.toxlet.2012.10.001
Because more than 95% human urinary samples have been detected to contain its metabolites (Meeker et al., 2007; Zhang et al., 2009), DEHP is believed to be ubiquitously present in human being. The United States Environmental Protection Agency (EPA) has classified DEHP as a chemical that may be potentially harmful to human health. Several studies indicate an association of phthalates with some potentially adverse endpoints, including interference with the timing of parturition, shortening anogenital distance of boys and lowering birth weight of neonates (Adibi et al., 2009; Swan, 2008; Zhang et al., 2006, 2009). In laboratory animals, DEHP has been reported to cause a dose-dependent increase in the incidence of Leydig cell tumors in adult Sprague Dawley rats when they were exposed to 0–300 mg/kg DEHP per day for 159 weeks (Voss et al., 2005). DEHP also induced a shortening of anogenital distance and lower testicular testosterone level of male pups in the postnatal day 2–3 when dams were exposed to 750 mg/kg DEHP from gestational day 14 to postnatal day 3 (Gray et al., 2000). Our previous study
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demonstrated that DEHP exposure during pubertal periods caused Leydig cell hyperplasia in rats (Akingbemi et al., 2001, 2004). Due to the proliferative capacity of Leydig cell precursors during puberty, increased mitosis of these precursor cells may be the cause of Leydig cell hyperplasia in rats (Akingbemi et al., 2001, 2004). However, there is still unclear whether DEHP exposure affects the Leydig cell functions at adulthood. In the adult testis, a small pool of stem cells regenerates to compensate the loss of mature Leydig cells. However, it is clear from studies of adult rats that the resident stem cells are able to support the restoration of adult populations of Leydig cells to testes from which the originally present cells have been experimentally eliminated by the alkylating agent ethane dimethane sulfonate (EDS) (Teerds, 1996). The loss of Leydig cells triggers a rapid regeneration of new adult population of Leydig cells within 4 weeks in the rat testis due to high circulatory level of luteinizing hormone (LH) (Kerr et al., 1985). Four days after the treatment of EDS, all Leydig cells are eliminated, and 14 days post-EDS newly formed progenitor Leydig cells appear and they gradually differentiate into more mature Leydig cells in this lineage from 21 to 35 days post-EDS (Teerds, 1996). In the present study, we reported the effects of DEHP exposures on the regeneration process of new population of Leydig cells at adulthood in EDS-treated model. 2. Materials and methods 2.1. Animal treatment The EDS was a gift from Dr. L. Earl Gray (U.S. Environmental Protection Agency, Research Triangle Park, NC). EDS was dissolved in a mixture of dimethyl sulfoxide (DMSO) and water (DMSO:H2 O, 1:4, v/v) for intraperitoneal injection. Twenty-four 90-day-old male Long-Evans rats (Charles River, Raleigh, NC) received a vehicle control (DMSO:H2 O, 1:4, v/v), and six rats per time-point were killed at days 4, 14, 21 and 35 post-EDS as the control. Seventy-two 90-day-old male Long-Evans rats (Charles River, Raleigh, NC) received a single intraperitoneal injection of 75 mg/kg EDS to eliminate Leydig cells, and then randomly divided into three groups with 24 rats that were gavaged with vehicle (corn oil 1.0 ml/250 g rat), 24 rats that were gavaged with 10 mg/kg/day DEHP (Sigma, CAS 204-211-0, purity of 99%) and 24 rats that were gavaged with 750 mg/kg/day DEHP. DEHP was gavaged 1 h after EDS for 35 days. DEHP was suspended in corn oil and gavaged to the rat by 0.5 ml/250 g rat. Dosages (0, 10, or 750 mg/kg per day DEHP) were selected based on our previously determined lowest-observed-effect level (10 mg/kg) of DEHP that increases Leydig cell number (Akingbemi et al., 2001; Ge et al., 2007), and lowest-observed-effect level (750 mg/kg) of DEHP that inhibits serum testosterone level in the pubertal rats (Foster et al., 2001; Ge et al., 2007). Eight rats per group were euthanized by CO2 on days 14, 21 and 35 post-EDS. Two hours before they were killed, all animals received an intraperitoneal injection of BrdU (40 g/g body weight, Cat. No. 280879 Roche Diagnostics) to label dividing cells. Blood samples were collected, placed in a gel tube and centrifuged at 1500 × g for 10 min to collect serum. The serum was stored at −20 ◦ C until the radioimmunoassay for T and LH. One testis from each animal was removed and frozen in liquid N2 for subsequent study of mRNA and steroidogenic enzyme levels, and the contralateral testis from each animal was punched holes using a needle and fixed by Bouin’s solution. All animal procedures were performed in accordance with the policies of The Rockefeller University’s Animal Care and Use Committee. 2.2. Serum testosterone and LH analysis Serum testosterone concentrations were measured with a tritium-based radioimmunoassay as previously described (Ge and Hardy, 1998). Interassay variation of the testosterone assay was between 7 and 8%. Serum LH concentrations were measured using the method described by Chandrashekar and Bartke (1988). LH standards and antibody were obtained through the National Hormone and Pituitary Program. Radioactive 125 I-rat LH was obtained through Covance Laboratories (Vienna, VA), and IgG antiserum was obtained from ICN Pharmaceuticals (Costa Mesa, CA). The sensitivity and inter- and intra-assay coefficients of variation for the LH assay were, respectively, 0.08 ng per ml and 5% and 6%.
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(Payne et al., 1980). After 3HSD reaction, nitro-blue tetrazoliumas is reduced and deposited as a diformazan at the site of Leydig cell cytosol. 4 ,6-Diamidino2-phenylindole (DAPI) was used as nuclear counterstaining of a cell. Testis sections were captured by using a Nikon Eclipse E800 microscope (Nikon) and a SPOT RT digital camera (model 2.3.0.; Diagnostic Instruments) interfaced to a computer. Cell numbers were estimated using image analysis software (Image-Pro Plus, Media Cybernetics, Rockville, MD). Leydig cell numbers were enumerated according to the fractionator technique (Akingbemi et al., 2004). Testes were cut and sections were collected every 15 section cuts for further analysis. About 10 sections were sampled randomly from each testis. The total number of Leydig cells per 100 seminiferous tubules was calculated since the tubule number was not affected by EDS. 2.4. BrdU incorporation Four testes per group per time-point were randomly selected. The testes were then removed, punched three holes per testis with needle to allow fixative to flow in and stored in the Bouin solution fixative for two days. After dehydration in ethanol and xylene, the testes were embedded in paraffin for immunocytochemical analysis. Leydig cells were identified through immunopositive staining for the marker enzyme 3HSD. In brief, 6-mm-thick transverse sections were prepared and mounted on glass slides (Cat. No. 12-550-15; Fisher Scientific Company, Hampton, NH). Avidin-biotin immunostaining was performed using a kit (Cat. Nos. PK-2200 for BrdU and PK-6101 for 3HSD; Vector Laboratories Inc., Burlingame, CA) according to the manufacturer’s instructions. Antigen retrieval was carried out by microwave irradiation for 10 min in 10 mM (pH 6.0) citrate buffer, and endogenous peroxidase was blocked with 0.5% H2 O2 in methanol for 30 min. The sections were then incubated with a monoclonal anti-BrdU antibody (RPN 202; Amersham Biosciences, Little Chalfont, UK) for 30 min at room temperature. The antibody bound to the nuclei was visualized with diaminobenzidine (Cat. No. sk-4100; Vector Laboratories Inc.) and the labeled nuclei were stained black by adding a nickel solution to the chromogen. After washing, the sections were double-labeled by incubation with a 3HSD polyclonal antibody diluted 1:3000 (provided by Dr. Van Luu-The, Laval University, Quebec, Canada) for 1 h at room temperature. The antibody–antigen complexes were visualized with diaminobenzidine alone, resulting in brown cytoplasmic staining in positively labeled Leydig cells. The sections were counterstained with Mayer hematoxylin, dehydrated in graded concentrations of alcohol, and cover-slipped with resin (Permount, SP15-100; Fisher Scientific). In control experiments, sections were incubated with nonimmune rabbit IgG (3HSD) or mouse IgG (BrdU) using the same working dilution as the primary antibody. 16 mosaic sections for each of three nonadjacent sections per testis were captured using a Nikon Eclipse E800 microscope (Nikon, Inc., Melville, NY) equipped with a 60 objective lens and a SPOT RT digital camera (model 2.3.0., Diagnostic Instruments, Inc., Michigan City, IN) interfaced to a computer. 200–300 Leydig cells were enumerated. Labeling index (LI) was calculated as BrdU positive cells per 100 Leydig cells. 2.5. Quantitative PCR Six testes per group per time-point were randomly selected. Total RNA was extracted from rat testes in Trizol according to the manufacturer’s instructions (Invitrogen, Carlsbad, CA). First-strand synthesis and real-time PCR (qPCR) were performed as previously described (Ge et al., 2005). Ribosomal protein S16 (Rps16) mRNA levels were assayed in all samples as an internal control. The expression levels of nine Leydig cell specific genes analyzed and the primers for the following genes were described in our previous study (Lin et al., 2008). These genes include receptor gene: luteinizing hormone receptor (Lhcgr); steroidogenic transporters and enzymes: scavenger receptor class B member 1 (Scarb1), steroidogenic acute regulatory protein (Star), cholesterol side chain cleavage enzyme (CYP11A1, encoded by Cyp11a1), 3HSD1 (Hsd3b1), 17␣-hydroxylase/20lyase (CYP17A1, encoded by Cyp17a1) and 17-hydroxysteroid dehydrogenase 3 (17HSD3, encoded by Hsd17b3); biomarker for Leydig cells at the more advanced stage: 11-hydroxysteroid dehydrogenase 1 (Hsd11b1) (Phillips et al., 1989); and Leydig cell differentiation status marker: insulin-like growth factor 3 (Insl3). 2.6. Homogenization and protein content assay The testes (randomly selected 6 samples per group per time-point) were homogenized in 1 ml ice-cold 0.1 M PBS (pH 7.2) containing 0.25 M sucrose. Supernatants were collected by centrifugation at 700 × g for 30 min. Supernatants were used to measure the protein level of CYP11A1, and enzyme activities of CYP17A1, 3HSD1 and 17HSD3. The protein concentrations were determined using a kit (No. 5000006, Bio-Rad Laboratories, Inc., Hercules, CA) with BSA as a standard. 2.7. Western blot analysis of CYP11A1
2.3. Determination of Leydig cell number Four testes per group per time-point were randomly selected. Frozen testis was sectioned in a cryostat, and 8 m testis sections were cut. Leydig cells were identified by the histochemical staining of Leydig cell specific marker 3-hydroxysteroid dehydrogenase (3HSD) activity with 0.4 mM etiocholanolone and 1 mg nitro-blue tetrazoliumas and NAD+ as cofactor according to the previously described method
Protein (50 g) was boiled in equal volumes of sample loading buffer, a Tris–Cl buffer (pH 6.8) containing 20% glycerol, 5% SDS, 3.1% dithiothreitol, and 0.001% bromophenol blue. Protein samples were electrophoresed on 10% polyacrylamide gels containing SDS. Proteins were electrophoretically transferred onto nitrocellulose membranes, and after 30 min exposure to 10% nonfat milk to block nonspecific binding, the membrane was incubated with a CYP11A1 antibody (catalogue no.
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RDIp450sccabr, RDI Research Diagnostics, Inc., Flanders, NJ; 1:1000). The membranes were then washed and incubated with a 1:5000 dilution of second antibody that was conjugated to horseradish peroxidase (anti-rabbit Ig, HRP-linked whole antibody produced in donkey; Amersham Biosciences Corp., Piscataway, NJ). The washing step was repeated, and immunoreactive bands were visualized by chemiluminescence using a kit (ECL, Amersham, Arlington Heights, IL). Then the antibody in the membrane was stripped away and probed again by an antibody against -actin (ACTB, Sigma; no. A2228; 1:1000). The second anti-mouse Ig, HRP-linked whole antibody produced in sheep (Amersham Biosciences, Piscataway, NJ) was used and the ACTB band was visualized by chemiluminescence. Protein levels were measured by densitometry of the films and normalized to ACTB. 2.8. Enzyme assay Testosterone biosynthetic enzymes activities of CYP17A1, 3HSD1 and 17HSD3 were determined by thin layer chromatography (TLC) and radiometry as previously described (Ye et al., 2011). The reaction mixtures (total volume of 250 l) containing 25–160 g protein, 0.2 mM cofactors (NAD+ for 3HSD1, NADPH for CYP17A1, and 17HSD3), and 1000 nM steroid substrates (radiolabeled plus cold substrates) were incubated in shaking water bath at 34 C for 1–3 h. The substrates were follows: pregnenolone (for 3HSD1), progesterone (for CYP17A1), and androstenedione (for 17HSD3). The preliminary experiment was conducted to determine the linear reaction curve using different concentrations of proteins at different time periods. The steroids were extracted from reaction mixture with 1 ml of ice-cold ether, and the organic layer was evaporated under nitrogen gas. The extract of steroids was suspended in 70 l ether and then spotted on thin layer plates (Baker-flex, Phillipsburg, NJ). The steroids were separated chromatographically in chloroform:methanol (97:3, v/v) for 3HSD1, and 17HSD3 as well as chloroform:ether (7:1, v/v) for CYP17A1. The radioactivity was measured with a scanning radiometer (System 200/AC3000, Bioscan, Inc., Washington DC). The conversion of steroid to product was calculated as a percentage of the total radioactivity found in the product. All assays were repeated in triplicate. 2.9. Statistical analysis Values are expressed as mean ± SEM, and data were analyzed using one-way ANOVA with Tukey’s comparison of all groups at each time point post-EDS using GraphPad Prism (version 5, GraphPad Software Inc., San Diego, CA). Significant difference was regarded at P < 0.05.
3. Results 3.1. Gestational exposure 3.1.1. General toxicological parameters and serum testosterone and LH levels Gavage of rats with 10 and 750 mg/kg DEHP for 35 days did not affect body weights (data not shown). After DEHP treatment, all rats were survived and had normal clinical signs, confirming the results of our previous study. Serum testosterone level was used as an indicator of the regeneration process. The baseline of testosterone level in the adult Long Evens male rats without EDS was about 5 ng/ml (Akingbemi et al., 2001). Serum testosterone levels were significantly reduced by 10 and 750 mg/kg DEHP on day 35 post-EDS (Fig. 1A). Serum LH level was significantly higher in the 10 mg/kg DEHP group on day 21 post-EDS. 3.1.2. Effects of DEHP on Leydig cell number and proliferation during regeneration Leydig cells can be identified by 3HSD staining (blue diformazan deposit). 3HSD staining reflects the Leydig cells but the status of Leydig cell differentiation. As shown in Fig. 2A, there were many 3HSDpos cells in the interstitium of 90-day-old rat testis. Four days post-EDS, all 3HSDpos cells were eliminated (Fig. 2B). 14 days post-EDS, there was one or 2 newly formed 3HSDpos cells in the control testis (Fig. 2C), while there were increased numbers of 3HSDpos cells in the DEHP-treated testes (Fig. 2D illustrates the 3HSDpos cells in the 10 mg/kg DEHP-treated testis). Enumeration of 3HSDpos cells showed that there were significantly and dose-dependently increased numbers of 3HSDpos cells in both DEHP-treated groups on day 14 (Fig. 3A), 21 (Fig. 3B) and 35 (Fig. 3C) post-EDS. The numbers of 3HSDpos cells in 10 and 750 mg/kg DEHP
Fig. 1. Serum testosterone and LH levels after DEHP exposure during the course of regeneration. Rats were i.p. treated 75 mg/kg EDS and 1 h later they were gavaged with 0 or 10 or 750 mg DEHP. Panel A: testosterone, and Panel B: LH. Bar , and represents 0 (control), 10 and 750 mg/kg DEHP, respectively. Mean ± SEM, n = 6, *significant change compared to control at each time point.
testes compared to control were increased by 3.5 and 9 fold on day 14. However, the number of 3HSDpos cells in both DEHP-treated groups was only about 1.36 fold of control on day 35 post-EDS. This indicates that DEHP treatment promotes the proliferation of 3HSDpos cells during earlier stage of regeneration. We further investigated the proliferation of Leydig cells using BrdU incorporation. As shown in Fig. 4, 10 and 750 mg/kg DEHP significantly increased the labeling index of Leydig precursor cells during regeneration on days 14 and 21 post-EDS. However, labeling index of Leydig cells never reached significance of both DEHPtreated groups when compared to the vehicle control on day 35 post-EDS (Fig. 4C). This indicates that the increased numbers of Leydig cells are mainly from the Leydig precursor cells at the earlier stage of regeneration. 3.1.3. Effects of DEHP on Leydig cell differentiation during regeneration Although there was the increased number of Leydig cells during the regeneration by DEHP, serum testosterone levels in both DEHP-treated groups were only half of that of control on day 35 post-EDS, indicating that there is defect of Leydig cell
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Fig. 2. Histochemical staining of 3HSD1 of testis. 3HSD1 was stained blue, and nucleus was fluorescently stained white by DAPI. Rats were i.p. treated 75 mg/kg EDS and 1 h later they were gavaged with 0 or 10 or 750 mg DEHP. Panel A: normal 94-day-old testis; Panel B: 14-day-post-EDS control testis; Panel C: 10 mg/kg DEHP-treated testis on day 14 post-EDS; and Panel D: 750 mg/kg DEHP-treated testis on day 14 post-EDS. Black arrows point to 3HSD1 positive cells. Bar = 50 m. (For interpretation of the references to color in Figure legend, the reader is referred to the web version of the article.)
differentiation in DEHP-treated testis. The expression levels of several Leydig cell-specific genes as biomarkers of differentiated status of Leydig cells during regeneration were shown in Fig. 5. We found that although Leydig cell numbers were increased by 3.5 and 9 fold in 10 and 750 mg/kg DEHP-treated testes on day 14 post-EDS, the expression levels of all selected Leydig cell specific genes were not altered. On day 21 post-EDS, only Star reached significant increase in 10 mg/kg DEHP group, while Insl3, a gene that reflects the differentiation status of Leydig cells and is not regulated by LH,
Leydig cell #/100 tubules
A. 14 days 600
was significantly decreased in both 10 and 750 mg/kg DEHP-treated testes. At that time point, the expression levels of steroidogenic enzyme genes Cyp11a1 and Hsd3b1 were significantly decreased in 750 mg/kg DEHP-treated testes. On day 35 post-EDS, the expression levels of Lhcgr, Cyp11a1, Hsd3b1 and Hsd11b1 were down-regulated in both 10 and 750 mg/kg DEHP-treated testes. We determined the protein levels of CYP11A1 and enzyme activities of CYP17A1, 3HSD1 and 17HSD3 on day 35 post-EDS, and found that CYP11A1 level and 3HSD1 activity were significantly
B. 21 days
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C. 35 days
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Fig. 3. Leydig cell number. Leydig cell numbers (#) were calculated as cells per 100 tubules. Mean ± SEM (n = 4). Rats were i.p. treated 75 mg/kg EDS and 1 hr later they were gavaged with 0 or 10 or 750 mg DEHP. Panel A: 14 days; Panel B: 21 days; and Panel C: 35 days post-EDS. *, **Significant differences compared to control at P < 0.05 and 0.01 at each time point post-EDS, respectively.
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Labeling index (100%)
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Fig. 4. Labeling index of Leydig cells. Labeling index was calculated as BrdU positive cells per 100 Leydig cells. Rats were i.p. treated 75 mg/kg EDS and 1 h later they were gavaged with 0 or 10 or 750 mg DEHP. Mean ± SEM (n = 4). Panel A: 14 days; Panel B: 21 days; and Panel C: 35 days post-EDS. Labeling index was calculated by the percentage of BrDU-positive cells per 200 interstitial cells. *, **Significant differences compared to control at P < 0.05 and 0.01 at each time point post-EDS, respectively.
reduced in both doses of DEHP while CYP17A1 and 17HSD3 were without changes (Fig. 6), confirming their mRNA level changes. 4. Discussion The EDS-treated rat Leydig cell regeneration model is very unique for observing the proliferation and differentiation of Leydig cells in the adult testis. Leydig cells in the adult testis are completely eliminated after a single intraperitoneal injection of EDS (Teerds, 1996). The loss of Leydig cells leads to the elevation of circulating LH, which together with local growth factors or cytokines promotes the proliferation of Leydig precursor cells and thereafter differentiation. The present study demonstrated that DEHP exposure to adult rats led to the increased Leydig cell number due to the increased the proliferative capacity of these newly formed Leydig precursor cells during the earlier regeneration process (from days 14 to 21 postEDS). Stereological enumeration of Leydig (3HSDpos ) cell numbers showed that the earlier exposure to DEHP have caused 3.5 and 9 fold on day 14, 1.5 and 1.8 fold on day 21, as well as 1.36 and 1.36 fold increase in Leydig cell numbers on day 35 post-EDS (Fig. 3). Apparently, the increased number of Leydig cells came from the proliferation of Leydig cell precursors from day 14 to 21 post-EDS (Fig. 4). The lack of significance for BrDU labeling at day 35 postEDS in not merely be a representation that spatial capacity of the interstitial compartment was reached The regeneration of Leydig cells showed a very similar pattern of pubertal Leydig cell development with higher proliferative rate in progenitor Leydig cells and lower proliferative rate later on (Ge and Hardy, 1997). The testis weight post-EDS treatment cannot reflect the Leydig cells because the testis weight was significantly reduced after EDS due to blocked spermatogenesis after loss of testosterone. Quite interestingly, high dose of DEHP exposure to adult rats inhibited the differentiation of Leydig cells, as judged by the decreased expression levels of Cyp11a1, Hsd3b1, Lhcgr, and Insl3, starting on day 21 post-EDS. Interestingly, Insl3, a gene that reflects the differentiation status of Leydig cells or Leydig cell numbers and is not regulated by LH, was also significantly decreased in low dose (10 mg/kg) and high dose (750 mg/kg) DEHP-treated testes. Since the Leydig cell numbers were actually increased in both doses of DEHP, the decreased expression level of Insl3 may reflect the delayed status of Leydig cell differentiation at 21 days post-EDS. Even at 35-day post EDS, the Leydig cell numbers were doubled in both doses of DEHP, the reduction of Insl3 expression levels in these groups indicates that the delayed differentiation of Leydig cells even at 35 days post-EDS. The serum level of testosterone
usually reflects the differentiated status of Leydig cells or the Leydig cell numbers in the testis. Apparently, this level reached significant decrease on day 35 post-EDS (Fig. 1A). There were no significant changes of serum testosterone levels from days 14 to 21 post-EDS, this might be due to significant higher ratio of number of Leydig precursor cells on days 14 (3.5–9 fold) and 21 (1.5 to 1.8 fold). The expression levels of Lhcgr, Hsd3b1 and Hsd11b1 in the DEHP-treated testes were decreased, even the Leydig cell numbers were increased, indicating that these Leydig cells may be less differentiated and produce fewer amount of testosterone. Other researchers also reported that the decreases in serum testosterone at high doses of DEHP when administered from weaning through adulthood without EDS administration (Noriega et al., 2009). The expression levels of Leydig cell specific genes, including Cyp11a1, Hsd3b1 were actually decreased though such higher ratio of Leydig cell numbers in high-dose DEHP-treated groups vs. the control group. The lowest observed adverse effect level (LOAEL) for increasing Leydig cell number and decreasing the differentiation of Leydig cells was 10 mg/kg/day. This level is within the suggested environmental exposure levels of the human population. Previously, our laboratory and others have demonstrated that low dose of DEHP can increase Leydig cell numbers during puberty by increasing the proliferation of Leydig precursor cells (Akingbemi et al., 2001, 2004). It is true that during puberty Leydig cell precursor cells (progenitor and immature Leydig cells) are proliferative (Ge and Hardy, 1998). Specifically, progenitor Leydig cells have higher proliferative capacity. However, at adulthood, cell replication slowed abruptly after puberty. The estimated turnover time for adult Leydig cells exceeds the two-year lifespan of the average rat (Kerr et al., 1987). In the present study, we also observed a significant increase of proliferation of Leydig cells during the earlier regeneration process in EDS-treated model, thus contributing to a significantly increased Leydig cell numbers. A recent study also showed that high dose of dibutyl phthalate, another widely used phthalate, significantly increased Leydig cell numbers during regeneration after EDS (Heng et al., 2011). This indicates that dibutyl phthalate and DEHP have a similar mechanism. However, the mechanism of stimulating Leydig cell proliferation in the regeneration model may be different from that caused by DEHP during pubertal exposure. Our previous study observed a significant increase LH levels by DEHP during pubertal exposure (Akingbemi et al., 2001, 2004). LH level is controlled by the circulating androgen. The normal adult Long Evens rats had serum LH levels around 0.5 ng/ml (Akingbemi et al., 2001). Loss of Leydig cells after EDS treatment significantly increased circulating LH level, and at 14 days post-EDS serum LH levels were still around 10 ng/ml
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Fig. 5. Steady-state mRNA levels of Leydig cell specific genes after DEHP treatment by quantitative PCR. Rats were i.p. treated 75 mg/kg EDS and 1 h later they were gavaged with 0 or 10 or 750 mg DEHP. Mean ± SEM (n = 5). (a)–(c) indicate significant differences of each DEHP-treated group compared to control (0 mg/kg DEHP) at each time point post-EDS at P < 0.05, 0.01, 0.001, respectively.
(Fig. 1B). In the present study, we only observed the increased LH level in the 10 mg/kg DEHP group on day 21 post-EDS, although there were significant increases of Leydig cell proliferation in both 10 and 750 mg/kg DEHP-treated groups. This difference may be complicated by the elevated LH levels in EDS-treated rats due to the negative feedback after elimination of Leydig cells. The change of LH levels may not be important factor to contribute to the proliferation of Leydig cell precursors, since the initial proliferation and
differentiation of Leydig cell precursors can occur in the absence of LH in hypogonadal rats (Teerds et al., 1989). In conclusion, the present study shows that DEHP exposure at adulthood increased Leydig cell numbers but inhibited the differentiation process during the regeneration. Leydig cell hyperplasia has been documented in rat neonates following in utero exposure to DEHP (Lin et al., 2008; Mahood et al., 2005; Mylchreest et al., 2002; Parks et al., 2000).
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Fig. 6. Protein level and steroidogenic enzyme activity of testis after DEHP treatment. CYP11A1 was measured by Western blotting (Panel A and B). Rats were i.p. treated 75 mg/kg EDS and 1 h later they were gavaged with 0 or 10 or 750 mg DEHP. 3HSD1 was measured as described as in Section 2. Mean ± SEM (n = 5–6). **, ***Significant difference of each DEHP-treated group compared to control (0 mg/kg DEHP) at P < 0.01 and 0.001, respectively.
Conflict of interest statement None. Acknowledgements We thank Chantal M. Sottas for the technical support from the Population Council. This work was also partially funded by NSFC 30871434 (R.S.G.) and Wenzhou Science & Technology Funding Project H20090075 (X. W.L.). Partial data were presented in American Society Annual Meeting held in 2009. References Adibi, J.J., Hauser, R., Williams, P.L., Whyatt, R.M., Calafat, A.M., Nelson, H., Herrick, R., Swan, S.H., 2009. Maternal urinary metabolites of di-(2-ethylhexyl) phthalate
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