Biol Reprod
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

BOR - Papers in Press, published online ahead of print November 24, 2004.
Biol Reprod 2004, 10.1095/biolreprod.104.031583
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
72/3/745    most recent
biolreprod.104.031583v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Bhattacharya, N.
Right arrow Articles by Kim, K. H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Bhattacharya, N.
Right arrow Articles by Kim, K. H.
Agricola
Right arrow Articles by Bhattacharya, N.
Right arrow Articles by Kim, K. H.
BIOLOGY OF REPRODUCTION 72, 745–754 (2005)
DOI: 10.1095/biolreprod.104.031583
© 2005 by the Society for the Study of Reproduction, Inc.

Differential Effects of Phthalates on the Testis and the Liver1

Nandini Bhattacharya3, Jannette M. Dufour3, My-Nuong Vo3, Janice Okita4, Richard Okita4, and Kwan Hee Kim2,3

School of Molecular Biosciences3 College of Pharmacy,4 Center for Reproductive Biology, Washington State University, Pullman, Washington 99164


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Phthalates have been shown to elicit contrasting effects on the testis and the liver, causing testicular degeneration and promoting abnormal hepatocyte proliferation and carcinogenesis. In the present study, we compared the effects of phthalates on testicular and liver cells to better understand the mechanisms by which phthalates cause testicular degeneration. In vivo treatment of rats with di-(2-ethylhexyl) phthalate (DEHP) caused a threefold increase of germ cell apoptosis in the testis, whereas apoptosis was not changed significantly in livers from the same animals. Western blot analyses revealed that peroxisome proliferator-activated receptor (PPAR) {alpha} is equally abundant in the liver and the testis, whereas PPAR{gamma} and retinoic acid receptor (RAR) {alpha} are expressed more in the testis. To determine whether the principal metabolite of DEHP, mono-(2-ethylhexyl) phthalate (MEHP), or a strong peroxisome proliferator, 4-chloro-6(2,3-xylindino)-2-pyrimidinylthioacetic acid (Wy-14,643), have a differential effect in Sertoli and liver cells by altering the function of RAR{alpha} and PPARs, their nuclear trafficking patterns were compared in Sertoli and liver cells after treatment. Both MEHP and Wy-14,643 increased the nuclear localization of PPAR{alpha} and PPAR{gamma} in Sertoli cells, but they decreased the nuclear localization of RAR{alpha}, as previously shown. Both PPAR{alpha} and PPAR{gamma} were in the nucleus and cytoplasm of liver cells, but RAR{alpha} was predominant in the cytoplasm, regardless of the treatment. At the molecular level, MEHP and Wy-14,643 reduced the amount of phosphorylated mitogen-activated protein kinase (activated MAPK) in Sertoli cells. In comparison, both MEHP and Wy-14,643 increased phosphorylated MAPK in liver cells. These results suggest that phthalates may cause contrasting effects on the testis and the liver by differential activation of the MAPK pathway, RAR{alpha}, PPAR{alpha}, and PPAR{gamma} in these organs.

di-(2-ethylhexyl) phthalate, di-isonanoyl phthalate, environment, liver, mitogen activated protein kinase, p38, peroxisome proliferator activated receptor, Sertoli cells, testis, toxicology


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Phthalic acid esters, such as di-(2-ethylhexyl) phthalate (DEHP) and di-isononyl phthalate (DINP), are used as plasticizers in commercial products such as plastic food wraps, children's toys, blood transfusion and dialysis bags, and catheters [1, 2]. However, phthalates are not covalently bound to the plastic material and leach into the environment. The potential for significant exposure to humans, especially to a sensitive pediatric population, is of particular concern [3, 4]. Once ingested, DEHP is hydrolyzed by intestinal esterases to mono-(2-ethylhexyl) phthalate (MEHP), the active testicular toxicant [5].

Phthalates are members of a structurally diverse group of chemicals known as peroxisome proliferators (PPs), which include hypolipidemic drugs such as 4-chloro-6(2,3-xylindino)-2-pyrimidinylthioacetic acid (Wy-14,643) and clofibrate, industrial solvents, and herbicides in addition to plasticizers [6]. Treatment with the most abundantly produced phthalate, DEHP, promotes abnormal proliferation, leading to hepatocarcinogenesis [6], and causes a seemingly opposite effect in the testis (degeneration of germ cells). More specifically, DEHP causes considerable damage to the somatic cells of the seminiferous tubules, Sertoli cells, and shedding of spermatocytes and spermatids, and eventually, it decreases sperm production [7]. Evidence indicates that Sertoli cells are one of the primary targets of phthalates in imparting testicular toxicity [8–10]. Leydig cells also are targeted in the testis by phthalates [11, 12].

The effects of PPs are postulated to be mediated primarily through the activation of peroxisome proliferator-activated receptors (PPARs), of which three subtypes exist: {alpha}, ß, and {gamma} [13]. These receptors form heterodimers with retinoid X receptors (RXRs), and they regulate the transcription of genes that contain PP-responsive elements. The PPs appear to exert their effects on tissues mainly by activation of PPAR{alpha} [6], which induces the expression of enzymes typically involved in fatty acid oxidation, including peroxisomal fatty acyl coenzyme A oxidase and microsomal cytochrome P4504A1 (CYP4A1) [14, 15]. This idea is supported by the phenotype of transgenic mice that lack PPAR{alpha}, which exhibit lipid droplet accumulation in their livers [16].

In the testis, PPAR{alpha} was shown to be responsible, at least in part, for the testicular damage seen with phthalate treatment [17, 18]. However, different from the tumorigenic effect on the liver, which was abolished in the PPAR{alpha}-null mice [17, 19, 20], the damage imparted by DEHP treatment, albeit at a lower level than that seen in the wild-type mice, remained in the PPAR{alpha}-null mice [17]. This result suggests that both PPAR{alpha}-dependent and PPAR{alpha}-independent mechanisms must be effective to account for the DEHP-mediated toxicity in the testis; thus, other PPARs or nonreceptor mechanisms also may participate in the testicular toxicity. Consistently, a recent study demonstrated that MEHP also can stimulate the transcriptional activity of PPAR{gamma} in both mice and humans [21], providing evidence that MEHP could act through PPAR{gamma}. Which PPAR isoform is responsible for the action of PPs in the testis, however, is yet to be determined.

Furthermore, in the testis, a counter-relationship exists between retinoic acid and PP signaling in Sertoli cells. The nuclear localization of retinoic acid receptor (RAR) {alpha} is inhibited by PPs in Sertoli cells, whereas PPAR{alpha} appears to distribute into the nucleus after PP treatment [22]. It was suggested that PPs may impair testicular function by inhibiting RAR{alpha} signaling [22], which is critical to normal testicular function [23]. However, to our knowledge, whether this counter-relationship between RAR{alpha} and PPARs exists in liver cells (or could be extended to PPAR{gamma} in the testis) remains to be investigated.

In addition, mitogen-activated protein kinase (MAPK) can regulate the nuclear localization of RAR{alpha} [24]. Whether Sertoli cells respond to PPs by activating MAPK signaling pathways is not known. However, PPs have been shown to impart growth-regulatory activities by activating MAPK signaling pathways in rodent liver [25]. Specifically, phosphorylation of MAPK (also referred to as extracellular signal-regulated kinase [ERK] 1 and 2) and p38 MAPK in primary hepatocytes and murine immortalized liver cell lines were reported to increase with PP treatments [26, 27]. In cultured rat primary hepatocytes, MAPK activation is accompanied by increased expression of immediate early genes, such as c-myc, c-jun, c-fos, junB, and egr-1 [25, 26, 28, 29]. Such growth-regulatory effects could account for the carcinogenic properties of PPs in the liver [27, 29].

The aim of the present study was to compare the effects of the phthalates, DEHP and DINP, which is a weaker PP than DEHP [30], on testes and livers from the same animals with those from age-matched littermates administered the solvent corn oil. Moreover, cultured Sertoli and liver cells were used to compare the molecular responses to MEHP and Wy-14,643 to gain insights regarding the mechanisms responsible for the eventual contrasting physiological effects on the testis and the liver. We show that both the testicular and liver cells respond rapidly to PP treatment by differential modulation of MAPK signaling pathways.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Chemicals

All-trans-retinoic acid (tRA) and Wy-14,643 were purchased from Sigma Chemical Co. (St. Louis, MO). The DEHP and DINP were purchased from Aldrich Chemical Co. (Milwaukee, WI). The MEHP was synthesized and purified by high-performance liquid chromatography to 99%, as determined by nuclear magnetic resonance and mass spectroscopy. The DEHP and DINP were dissolved in Super Refined Corn Oil (Croda, Inc., Parsippany, NJ) for gavaging rats. Other chemicals were dissolved in 0.1% (v/v) dimethyl sulfoxide (DMSO) and carried to the cell membrane by 0.1% fatty acid-free BSA (Boehringer Mannheim, Indianapolis, IN). Bis[amino[(2-aminophenyl)thio]methylene] (U0126; Biomol Research Lab, Plymouth Meeting, PA) is an inhibitor for MAPK kinase (Mek1 and Mek2), which is upstream of the ERKs in the MAPK signaling pathway, with a median inhibitory concentration of 72 nM for Mek1 and 58 nM for Mek2. This specific MAPK inhibitor has negligible effects on the kinase activities of protein kinase C, Abl, Raf, Erk, and c-Jun-N-terminal kinase [31]. It was dissolved in DMSO as recommended and used at a final concentration of 10 µM for treating primary Sertoli cells.

Animals

Male Sprague-Dawley rats were obtained from an in-house vivarium, fed ad libitum with Harlan Teklad 22/5 Rodent diet W (Harlan Teklad, Madison, WI), and treated at 30 days of age. At least three rats per group were treated with a single dose of DEHP (1200 mg/kg) or DINP (1200 mg/kg) in Super Refined Corn Oil by gavage and then killed at 0, 3, 6, 9, 12, 16, and 24 h. The DEHP was administered to rats instead of MEHP, because DEHP is the form that animals most likely are exposed to in the environment. Control rats received a similar volume of corn oil (1 ml/kg) and were killed at the same timepoints. For comparison, untreated rats of the same age were used as additional controls. Testes and livers were collected from each rat, fixed in Bouin for 6 h, embedded in paraffin, sectioned (thickness, 3 µm), and mounted onto 2% aminopropyltriethoxysilane-coated slides (Sigma). Testes and livers from at least three rats treated with DEHP also were processed to obtain protein extracts and RNA. Animal experimentation was approved by the Washington State University Institutional Animal Care and Use Committee and conducted in accordance with the highest standards of humane care as outlined in the NIH Guide for the Care and Use of Laboratory Animals.

Cell Cultures

A mouse Sertoli cell line, MSC-1 [32], was maintained in Dulbecco modified Eagle medium (DMEM; Invitrogen Life Technologies, Carlsbad, CA) containing 5% fetal calf serum (FCS) supplemented with penicillin (105 IU/L) and streptomycin (100 mg/L) at 37°C in a 5% CO2 atmosphere. Cells were grown to approximately 50% confluency before being serum-starved for 24–48 h with 0.1% FCS in DMEM to reduce endogenous tRA. The cells were treated with 0.5 mM Wy-14,643 or MEHP in media containing 0.1% FCS, 5 mM Hepes (pH 7.1), and 0.1% DMSO as the diluent. Primary Sertoli cells were isolated from the testes of 20-day-old rats by sequential enzymatic digestion as described previously [33]. Decapsulated testis fragments were digested with 0.25% (w/v) trypsin (Invitrogen) followed by 0.7 mg/ml of collagenase (Sigma) and 1 mg/ml of hyaluronidase (Sigma). Sertoli cells were plated under serum-free conditions on 100-mm plates (Nalge Nunc International, Rochester, NY) and were maintained in Ham F-12 medium (Invitrogen) at 32°C in a 5% CO2 atmosphere for a maximum of 5 days. After culture for 2 days, medium was changed every day to reduce endogenous tRA. The percentage of Sertoli cells was determined to be approximately 90% [22].

An immortalized cell line derived from a CD-1 mouse liver tumor that is nontumorigenic in nude mice [26], ML457 was maintained in DMEM supplemented with 10% FCS and 50 µg/ml of gentamicin (Invitrogen) at 37°C in a 5% CO2 atmosphere. Before stimulation, confluent cultures were serum-starved in 0.1% FCS containing medium for 3 days following the previous protocol [26]. The quiescent cells were then subjected to 0.5 or 1 mM Wy-14,643 or MEHP diluted in media containing 0.1% serum and 5 mM Hepes (pH 7.1) for the specified timepoints as described previously [26].

TUNEL Assay

The TUNEL assay (Apoptosis Detection System; Promega Biotech Corporation, Madison, WI) was conducted according to the manufacturer's instructions. This assay detects fragmented DNA in apoptotic cells by catalytic incorporation of fluorescein-12-dUTP at the 3'-OH ends of DNA using the terminal deoxynucleotidyl transferase (TdT) enzyme. A positive reaction was characterized by bright-green fluorescence at the site of the fluorescein 12-dUTP incorporation and was visualized directly using a Leitz DMRB microscope (Leica, Wetzlar, Germany) with epifluorescence and recorded by a Magnafire digital camera (Optronics, Goleta, CA). Sections incubated without the TdT enzyme served as negative controls for the assay. A positive control was pretreated with Deoxyribonuclease I, which resulted in multiple DNA fragments where fluorescein-dUTP was incorporated.

To quantitate the relative differences in the number of apoptotic cells between the control and treated testes, seminiferous tubules with greater than three or more TUNEL-positive cells out of 200 seminiferous tubules were counted for at least three rats. This quantitation procedure was adopted from a previous report [34, 35]. To quantitate the percentage of TUNEL-positive hepatocytes, the approximate number of hepatocytes in a defined area (12 squares) was counted (1100 hepatocytes/12 squares). Then, the percentage of TUNEL-positive cells was determined by counting the number of TUNEL-positive cells in the same area, dividing by 1100, and multiplying by 100. The TUNEL analyses were carried out on sections from at least three rats.

RNA Isolation and Reverse Transcriptase-Polymerase Chain Reaction

Total RNA from testis and liver tissues was processed according to the single-step method of Chomczynski and Sacchi [36] with minor alterations. The RNA from primary Sertoli cells was isolated using an RNAqueous kit (Ambion, Austin, TX). The purity and concentration of RNA was measured by ultraviolet (UV) spectrometry.

The cDNA was synthesized using Superscript II reverse transcriptase (RT; Invitrogen). Briefly, 1 µg of RNA, 0.5 µg of oligo-dT primer (Invitrogen), and 0.5 mM dNTPs were used in a final volume of 20 µl. To assess the degree of contaminating genomic DNA, parallel reactions for each RNA sample were run in the absence of the RT. The polymerase chain reaction (PCR) contained 2 µl of cDNA, 0.1 volume of 10x DNA polymerase buffer (200 mM Tris-HCl [pH 8.4], 500 mM KCl), 0.2 µM 5' and 3' primers, 0.2 mM dNTPs, 50 mM MgCl2, and Taq Polymerase (Promega) in a final volume of 50 µl. Primers were designed using the Primer Express software (Version 2; Applied Biosystems, Foster City, CA). The primers were as follows: PPAR{alpha}, TTCGGAAACTGCAGACCT and TTAGGAACTCTCGGGTGAT (442 base pairs [bp]) [37] or TGGAGTCCACGCATCTGAAG and CCGAATAGTTCGCCGAAAGA (51 bp); PPARß, CATCCTCACTGGCAAGTGCAG and GCAGCGGTAGAACACATGCA (155 bp); PPAR{gamma}, ACTCCCATTCCTTTGACATC and TCCCCACAGACTCGGCACTC (265 bp); and ribosomal S2 protein, CTGCTCCTGTGCCCAAGAAG and AAGGTGGCCTTGGCAAAGTT (105 bp). The PCR reaction was conducted in Eppendorf Mastercycler (Brinkmann Instruments, Westbury, NY) with the following program: denaturation for 30 sec at 95°C, annealing for 1 min at 55°C, and extension for 1 min at 72°C for 30 cycles. The PCR products were resolved on a 2: 1 GenePure LE Agarose (BioExpress, Kaysville, UT) or NuSieve GTG Agarose (FMC Bioproducts, Rockland, ME), stained with ethidium bromide, visualized by UV light source, and photographed with a DC290 ZOOM Digital camera (Eastman Kodak Corporation, Rochester, NY).

Protein Extract Preparation and Western Blot Analysis

The cells and tissues were washed with cold PBS and lysed in lysis buffer (5 mM EDTA, 1 mM EGTA, 50 mM Tris-HCl [pH 7.5], 250 mM NaCl, 0.1% Triton X-100, 50 mM NaF) containing Complete Mini Protease Inhibitor Cocktail Tablets (Roche Applied Science, Indianapolis, IN) at the recommended concentrations (100 µM PMSF, 10 µM aprotinin, 10 µM leupeptin). The lysates were incubated on ice for 30 min and then centrifuged for 30 min at 10 000 rpm. The supernatant was collected, and the protein concentration was determined [38] with BSA as a standard.

Twenty micrograms of the testicular and liver lysates were subjected to electrophoresis on the same 10% SDS-polyacrylamide gel and transferred to Immobilon-P membranes (Millipore, Bedford, MA). Membranes were blocked with 5% (w/v) Blotto (Carnation, Los Angeles, CA). The membranes were incubated with the following primary antibodies: antiphospho-p44/42 MAPK at a 1:400 dilution (Cell Signaling Technologies, Beverly, MA), antiphospho-p38 (Cell Signaling Technologies) at a 1:300 dilution, anti-ERK2 at a 1:400 dilution (Santa Cruz Biotechnology, Santa Cruz, CA), anti-CYP4A1 at a 1:150 dilution [39], anti-RAR{alpha} at a 1:300 dilution (Santa Cruz Biotechnology), anti-PPAR{alpha} at a 1:300 dilution (Santa Cruz Biotechnology), and anti-PPAR{gamma} at a 1:500 dilution (Biomol). Then, membranes were further incubated with secondary horseradish peroxidase-conjugated anti-rabbit immunoglobulin G antibody at a 1:10 000 dilution (ICN Biomedicals, Aurora, OH), and antibody-antigen complexes were detected by the Enhanced Chemiluminescence System (Amersham Pharmacia Biotech, Piscataway, NJ). After chemiluminescence detection, the same membranes were stained with Coomassie blue to evaluate the amount of proteins in each lane. Experiments were repeated at least three times.

Immunofluorescence

Both MSC-1 and ML-457 cells were plated in 24-well plates (Becton Dickinson Co., Franklin Lake, NJ) on 13-mm Thermanox plastic coverslips (Nalge Nunc) and fixed with methanol for 20 min at –20°C. Cells were blocked with 10% goat serum for 2 h before overnight incubation with a 1:300 dilution of anti-RAR{alpha}, anti-PPAR{alpha}, anti-RXR{alpha} (Santa Cruz Biotechnology) or a 1:500 dilution of anti-PPAR{gamma} (Biomol) at 4°C. Cells were washed three times with 1x PBS and incubated with a biotinylated secondary antibody at a 1:300 dilution for 1 h. Detection of antigen-antibody complexes was conducted using fluorescein avidin D (Vector Laboratories, Burlingame, CA). For a negative control, the RAR{alpha} and PPAR{alpha} antibodies were incubated with the immunizing peptides (Santa Cruz Biotechnology). For negative controls for PPAR{gamma} and RXR{alpha} antibodies, the cells were fixed, and further incubations were conducted without the addition of primary antibodies. The secondary antibody was omitted to detect background fluorescence as an additional control. The cells were counterstained with Vectashield (Vector Laboratories) containing 4',6'-diamidino-2-phenylindole or propidium iodide to visualize the nucleus. Images were obtained and digitized using a laser-scanning confocal system (MRC 1024; Bio-Rad, Hercules, CA).

Statistics

For statistical analysis, one-way ANOVA was performed, followed by pairwise comparisons of the means at P = 0.05 (Minitab 10 Xtra; Minitab, Inc., State College, PA).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
DEHP Treatment Promotes Apoptosis Within Testis But Not Liver

To study the extent of apoptosis after phthalate treatment, 30-day-old Sprague-Dawley rats were gavaged with DEHP or DINP, and the testis and liver were collected at 0, 3, 6, 9, 12, 16, and 24 h. Then, TUNEL assays were performed to detect DNA fragmentation on the tissue sections. In the testis, the fluorescent signals, indicative of apoptotic cells, were localized primarily to the germ cells very close to the basal side of the seminiferous tubules (Fig. 1, B and C). These cells were mostly primary spermatocytes, as determined by their position in the seminiferous tubules. The percentage of apoptotic tubules was obtained by counting the seminiferous tubules with greater than three fluorescent cells out of 200 tubules and then calculating the percentage. A threefold increase in apoptosis was observed after 12 h of DEHP treatment when compared to the Super Refined Corn Oil-treated controls (Fig. 1K). The levels returned to those of the corn oil-treated controls by 24 h. Morphologically, tubule sections from the treated rats exhibited increased sloughing of germ cells from the seminiferous epithelium into the lumen (Fig. 1G) and disorganization of germ cells (Fig. 1H) compared to the corn oil control sections (Fig. 1E). In contrast, DINP, a weaker peroxisome proliferator, did not significantly increase the number of TUNEL-positive cells in the testis (Fig. 1, J and K) or change the morphology of the testis (data not shown). Very few apoptotic cells were detected in the hepatocyte sections from DEHP-treated rats compared to the untreated hepatocyte sections (Fig. 2, compare A with B). No significant change was observed in the number of TUNEL-positive liver cells at any time after DEHP treatment (Fig. 2H). Additionally, liver cells from DEHP-treated rats including the sinusoid lining adjacent to the hepatocytes (Fig. 2, compare F and G) did not display any notable morphological changes compared to the control. Similarly, DINP did not significantly increase the number of apoptotic cells in the liver compared to the corn oil-treated control at any time after treatment (Fig. 2, compare D, E, and H).



View larger version (109K):
[in this window]
[in a new window]
  		FIG. 1. Morphology and TUNEL analysis of the testis after DEHP or DINP treatment. The TUNEL assay was performed on testicular sections from corn-oil treated 30-day-old rats (A and I), rats treated with DEHP for 12 h (B) and 16 h (C), and rats treated with DINP for 12 h (J). A TUNEL-positive cell is characterized by a bright green fluorescent dot. A positive-control section (D) was pretreated with DNase before TUNEL analysis. Representative testicular sections stained with hematoxylin-eosin from corn oil-treated control rats (E) and rats treated with DEHP for 24 h (F–H) are shown. White arrows indicate primary spermatocytes; a black triangle indicates vacuolization. The TUNEL-positive tubules were counted, and the results were plotted as the percentage of apoptotic tubules (K). Data are presented as the mean ± SD (n ≥ 3). An asterisk denotes a significant difference from the control level by ANOVA and Tukey honestly significant difference test (P < 0.05). White columns represent corn oil-treated controls, black columns DEHP treatment, and gray columns DINP treatment. Bar (A–F, I, J) = 50 µm as shown in D. Bar (G) = 50 µm as shown in H



View larger version (68K):
[in this window]
[in a new window]
  		FIG. 2. Morphology and TUNEL analysis of the liver after DEHP or DINP treatment. The TUNEL assay was performed on liver sections from corn oil-treated 30-day-old rats (A and D) and from rats 12 h after DEHP treatment (B) or DINP treatment (E). A positive-control section was pretreated with DNase (C). A representative liver section stained with hematoxylin-eosin from rats treated with corn-oil vehicle (F) or DEHP for 16 h (G) is shown. The TUNEL-positive liver cells were counted, and the percentage apoptotic cells were graphed (H). Data are presented as the mean ± SD (n ≥ 3). White columns, corn oil treated controls; black columns, DEHP treatment; gray columns, DINP treatment. The values for the treated samples were not significantly different from the corn oil-treated control samples by ANOVA and Tukey honestly significant difference test (P < 0.05). Bar = 100 µm (A–E) and 50 µm (F, G) as shown in F

Differential Expression of PPAR and RAR{alpha} in Testis Compared to Liver

The expression of PPAR{alpha}, PPAR{gamma}, and RAR{alpha} was evaluated in testes and livers isolated from 30-day-old rats treated with DEHP and killed at 3, 6, 12, 16, and 24 h after the treatment. Western blot analyses conducted on the protein lysates showed that the relative abundance of PPAR{alpha} protein was similar in the testis and the liver, whereas more PPAR{gamma} was found in the testis than in the liver (Fig. 3A). The RAR{alpha} was abundant in the testicular protein lysate, but it was barely detectable in the liver protein lysate (Fig. 3A). Conversely, CYP4A1 was detected in the liver lysate at 51 kDa but not in the testicular lysate (Fig. 3A). Interestingly, no extensive changes were apparent in the abundance of PPAR{alpha}, PPAR{gamma}, or RAR{alpha} in both the testis and the liver after DEHP treatment. In contrast, CYP4A1 increased considerably 12 h after DEHP treatment in the liver. The same blot used for Western blot analysis also was stained with Coomassie blue dye to verify even protein loading (Fig. 3B).



View larger version (64K):
[in this window]
[in a new window]
  		FIG. 3. Western blot analyses of PPAR{alpha}, PPAR{gamma}, RAR{alpha}, and CYP4A1 proteins in the testis and the liver. A) Thirty-day old rats were treated with DEHP, and cell extracts were collected from the testis (lanes 1–4) and the liver (lanes 5–8) from the same animals at 0 h (lanes 1 and 5), 12 h (lanes 2 and 6), 16 h (lanes 3 and 7) and 24 hr (lanes 4 and 8) after DEHP treatment. The PPAR{alpha}, PPAR{gamma}, RAR{alpha}, and CYP4A1 proteins were detected by Western blot analysis using their respective antibodies. B) Equivalent loading for each lane was verified by staining the same membranes with Coomassie blue dye. A representative blot from three separate experiments is shown

To determine whether the levels of PPARß mRNA in the testis and the liver are different from the levels of PPAR{alpha} and PPAR{gamma} mRNA, their transcripts were detected by RT-PCR analysis of RNA collected from the tissues at 12 and 24 h postgavaging. The S2 ribosomal protein served as an internal control and as a control to normalize between the levels of specific RT-PCR products from the testis and liver tissues. After DEHP treatment, PPAR{alpha}, PPARß, and PPAR{gamma} transcript levels did not change substantially in both the testis and the liver (Fig. 4, A and B). However, both the testis and the liver clearly contained substantial amounts of all PPAR transcripts. In the liver, PPARß transcript levels (Fig. 4B) were the highest, with moderate levels of PPAR{alpha} and PPAR{gamma}. The RT-PCR analysis also was conducted on RNA collected from primary Sertoli cells to confirm PPAR expression in Sertoli cells (Fig. 4C). Similar to the PPAR transcript levels in the testis, which is composed of many types of cells, including Sertoli cells, all PPAR transcripts were present in Sertoli cells.



View larger version (48K):
[in this window]
[in a new window]
  		FIG. 4. Transcript levels for PPAR{alpha}, PPARß, and PPAR{gamma} in the testis, liver, and primary Sertoli cells. Thirty-day old rats were treated with DEHP and RNA collected from the testis (A; lanes 1–3) and the liver (B; lanes 4 and 5) from the same animals at 0 h (lanes 1 and 4), 12 h (lanes 2 and 5), and 24 h (lanes 3 and 6) after DEHP treatment. The RNA also was isolated from testicular samples and primary Sertoli cells (C; lanes 7 and 8). The RT-PCR products representing PPAR{alpha}, PPARß, and PPAR{gamma} transcripts were separated on agarose gels. As an internal control, the same cDNA templates were amplified with primers to S2 ribosomal protein. The S2 levels were used to normalize the transcript levels of the receptors. The position of the products is shown on the left of each panel. Data are representatives of at least three experiments

Subcellular Localization of the Receptors after MEHP Treatment

Previously, we showed that MEHP inhibits tRA-induced nuclear localization of RAR{alpha} in primary Sertoli cells and MSC-1 cells; thus, RAR{alpha} was cytoplasmic in the presence of tRA and MEHP or tRA and Wy-14,643 [22]. Conversely, PPAR{alpha} translocated to the nucleus in the presence of MEHP or Wy-14,643 in primary Sertoli cells and MSC-1 cells [22]. To determine whether RAR{alpha} in liver cells responded to MEHP in a similar manner, ML-457 cells, an immortalized liver cell line, were treated tRA, MEHP, or both at the same concentrations as used previously. In contrast to its effect in Sertoli cells, tRA was not able to induce nuclear localization of RAR{alpha} in ML-457 cells. Under this circumstance, the distribution of RAR{alpha} was mainly cytoplasmic in ML-457 cells, regardless of the treatment (Fig. 5). In contrast, the distribution of PPAR{alpha} in ML-457 cells was mostly nuclear, but it also present in the cytoplasm (Fig. 5). This distribution did not change with treatment. Similarly, the distribution of RXR{alpha}, a potential partner for RARs and PPARs, was mainly nuclear. The RXR{alpha} distribution pattern did not alter with any of the treatments in either MSC-1 [22] or ML-457 cells (Fig. 5).



View larger version (84K):
[in this window]
[in a new window]
  		FIG. 5. The subcellular localization pattern of RAR{alpha}, PPAR{alpha}, and RXR{alpha} proteins in a liver cell line. The ML-457 cells were grown on coverslips and incubated for 60 min with vehicle (A, E, I, M, Q, and U), 1 µM tRA (B, F, J, N, R, and V), and 200 µM MEHP (C, G, K, O, S, and W). In addition, cells were pretreated with 200 µM MEHP for 30 min before incubation with both MEHP and tRA for an additional 60 min (D, H, L, P, T, and X). The cells were methanol-fixed and incubated with an anti-RAR{alpha} (A–D), anti-PPAR{alpha} (I–L), or anti-RXR{alpha} (Q–T) antibody followed by biotinylated goat anti-rabbit antibody and fluorescein avidin D (FITC) to locate receptors. The cells also were stained with 4',6-diamidino-2-phenylindole (DAPI; E–H and U–X) or propidium iodide (PI; M–P) to stain DNA in the nucleus. Bar =50 µm

When both MEHP and tRA were added to MSC-1 cells, however, the distribution of PPAR{gamma} was predominantly nuclear (Fig. 6), similar to that seen previously for PPAR{alpha} [22]. Treatment with MEHP alone resulted in concentration of PPAR{gamma} around the nuclear envelope in approximately 50% of MSC-1 cells (Fig. 6). On the other hand, in ML-457 liver cells, PPAR{gamma} showed a speckled nuclear distribution with some diffused staining in the cytoplasm, regardless of treatment with tRA, MEHP, or both (Fig. 6).



View larger version (122K):
[in this window]
[in a new window]
  		FIG. 6. Immunofluorescence microscopy demonstrating the subcellular localization pattern of PPAR{gamma} in Sertoli and liver cell lines. The MSC-1 (A–H) or liver ML-457 cells (I–P) were grown on coverslips and incubated for 60 min with vehicle (A, E, I, and M), 1µM tRA (B, F, J, and N), and 200 µM MEHP (C, G, K, and O). In addition, cells were pretreated with 200 µM MEHP for 30 min before incubation with both MEHP and tRA for an additional 60 min (D, H, L, and P). The cells were fixed in methanol and stained with an anti-PPAR{gamma} antibody, followed by biotinylated goat anti-rabbit antibody and fluorescein avidin D (FITC; A–D and I–L). Cells also were stained with 4',6-diamidino-2-phenylindole (DAPI; E–H and M–P) to stain DNA in the nucleus. Bar = 50 µm

MEHP Reduced Activation of MAPK in Sertoli Cells But Promoted Activation in Liver Cells

The amount of phosphorylated MAPK (ERK1 and ERK2) in MEHP-treated samples was detected with an antibody that recognized phosphorylated ERK1 and ERK2, and the levels were compared to those in the control samples treated with DMSO, the solvent used for MEHP. We found that DMSO induced the phosphorylation of ERK1 and ERK2 within 5 min (Fig. 7A, lanes 2 and 9) compared to the levels in the untreated control (Fig. 7A, lanes 1 and 8). This level of phosphorylation after DMSO treatment was maintained throughout the 20-min time period (Fig. 7A, lanes 2–4 and 9–11). However, MEHP inhibited the phosphorylation of ERK1 and ERK2 in primary rat Sertoli cells (Fig. 7A, lane 7) and in MSC-1 cells (data not shown). Similarly, a reduction of phosphorylated ERK1 and ERK2 in primary Sertoli cells was detected following treatment with another potent PP, Wy-14,643, for 5, 10, and 20 min (Fig. 7A, lanes 12–14). It appears that Wy-14,643 is more potent than MEHP (Fig. 7C). The amount of ERK1 and ERK2 in the protein lysates, as determined by an antibody that recognizes both phosphorylated and unphosphorylated ERKs, was equivalent in each lane (Fig. 7B). Additionally, the primary Sertoli cells were treated with an inhibitor to MAPK kinase (Mek), which is an upstream protein that phosphorylates ERKs in the MAPK signaling pathway. The pretreatment or cotreatment with an Mek inhibitor, U0296, further reduced the amount of phosphorylated ERK1 and ERK2 seen after MEHP and Wy-14,643 treatments in primary Sertoli cells (data not shown), indicating that PPs are acting similarly to U0296 as inhibitors of the MAPK signaling pathway in Sertoli cells.



View larger version (32K):
[in this window]
[in a new window]
  		FIG. 7. Effects of MEHP and Wy-14,643 on MAPK phosphorylation in rat primary Sertoli cells. The phosphorylation status of MAPK (A) was assessed on cellular extracts from primary rat Sertoli cells treated with DMSO vehicle (lanes 1–4 and 8–11), 0.5 mM MEHP (lanes 5–7), or Wy-14,643 (lanes 12–14) for 0, 5, 10, or 20 min using an antibody that recognizes phosphorylated ERK1 (p42) and ERK2 (p44). Equal protein loading was determined by reprobing the membrane with an anti-ERK2 antibody, which recognizes total ERK1 and ERK2 proteins (B). Phosphorylated and total MAPK proteins are indicated on the left. These experiments were conducted at least five times. The levels of phosphorylated MAPK (C) were determined by densitometric analysis of three independent experiments, normalized, and the relative fold-differences plotted (mean ± SD). Different value labels indicate a significant difference of means by ANOVA and Tukey honestly significant difference test (P < 0.05)

On the other hand, both MEHP and Wy-14,643 treatment for 20 min induced the phosphorylation of ERK1 and ERK2 in ML-457 liver cells (Fig. 8A, lanes 2–4 and 6–8) compared to the control DMSO-treated samples (Fig. 8A, lane 1). The phosphorylation of ERK1 and ERK2 was maximal at both 0.5 and 1 mM MEHP (Fig. 8A, lanes 3 and 4) or Wy-14,643 (Fig. 8A, lanes 7 and 8). The same liver protein samples also were probed with an antibody to ERK2, which recognizes total MAPK to verify an equal loading of proteins in each lane (Fig. 8B). Moreover, MEHP increased phosphorylated ERK1 and ERK2 in a time-dependent manner (Fig. 8C). Furthermore, phospho-p38 MAPK levels were induced in ML-457 cells following treatment with MEHP (Fig. 8D, lane 5) compared to the DMSO-treated controls (Fig. 8D, lane 4). However, in contrast to liver cells, incubation of primary rat Sertoli cells with either MEHP or Wy-14,643 did not alter the levels of phospho p38 MAPK (Fig. 8D, lanes 2 and 3).



View larger version (31K):
[in this window]
[in a new window]
  		FIG. 8. Effects of MEHP and Wy-14,643 on MAPK phosphorylation in ML-457 liver cells. The MAPK phosphorylation status was assessed on cell extracts from ML-457 liver cells treated with 0, 0.1, 0.5, and 1.0 mM MEHP (A; lanes 1–4) or Wy-14,643 (A; lanes 5–8) for 20 min and with 1.0 mM MEHP for 5, 10, and 20 min (C) using an anti-phospho-MAPK antibody. Additionally, phosphorylation of p38 was evaluated on cell lysates from primary Sertoli cells and ML-457 liver cells treated with 0.5 mM MEHP (lanes 2 and 5) or Wy-14,643 (lane 3) for 20 min (D). Equal protein loading was determined by reprobing the same membrane with an anti-ERK2 antibody (B and E). As controls, cells were treated with DMSO for 20 min (control, 0 mM). Experiments were conducted at least three times


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Differential control of gene expression is critical for the specification and maintenance of differentiated cell types that comprise various tissues. Environmental toxins could alter gene expression, and this may lead to abnormal proliferation, differentiation, and apoptosis in a variety of tissues. The purpose of the present study was to compare the molecular responses of the testis and the liver to a peroxisome proliferator, DEHP, and its principal metabolite, MEHP, to gain better insights regarding the mechanism of toxic action of phthalates on the testis. More specifically, the expression and activity of nuclear receptors such as RAR{alpha} and PPAR, which regulate transcription of their specific responsive genes, were examined to assess potential changes in gene expression that could accompany phthalate treatments of the testis.

Examination of the testis and the liver obtained from the same 30-day-old Sprague-Dawley rats treated with DEHP demonstrated that DEHP causes apoptosis in the testis but not in the liver. These results are similar to previously documented results obtained from Fischer rats [34] showing that spermatocytes undergo apoptosis as early as 6 h after MEHP exposure. The delay in time for the maximal apoptosis that we observed compared to that in the previous report [34] could be caused by the time that it takes for DEHP to be hydrolyzed to MEHP. Results from the liver also are within the realm of a previous report [40], which indicated a decrease in apoptosis, although in the present study, the decrease was not statistically significant. The same group further noted an induction of DNA synthesis in response to DEHP in the rodent liver [40].

Increased apoptosis in the testis was accompanied by morphological changes, such as obvious disorganization of germ cells, appearance of vacuolization, and detachment and sloughing of germ cells into the lumen of the tubule, that were detected as early as 6 h after DEHP treatment and became more severe by 24 h. The DEHP-induced apoptosis within the testis was observed prevalently in primary spermatocytes and was maximal by 12 h after DEHP treatment. The effect of DEHP on the testis is specific to DEHP, because a weaker peroxisome proliferator, DINP, did not significantly increase apoptosis in the testis. The DINP results are compatible with those of recent studies that deemed DINP to be less potent as a reproductive toxin compared to DEHP [3, 4].

Using RNA samples obtained from the testis and the liver of the same 30-day-old treated animals, we found that PPAR transcripts were as abundant in the testis as in the liver. More interestingly, PPs did not appreciably change the relative abundance of PPAR transcripts after DEHP exposure. In addition, primary Sertoli cells also expressed all PPAR transcripts. However, at the protein level, more PPAR{gamma} was detected in the testis than in the liver, whereas the relative abundance of PPAR{alpha} was similar in the testis and liver. Taken together, although these results can be construed as indicating that all three PPAR isoforms are available in the testis, including Sertoli cells, and in the liver, it seems to be clear that primary receptors functioning in the testis probably are different from those in the liver.

More specifically, it is interesting that the relative abundance of RAR{alpha} was high in the testis compared to the level in the liver. Previously, RAR{alpha} was demonstrated to be a critical receptor for testicular function [23] and to be expressed in both germ cells and Sertoli cells [41]. Moreover, we showed that lack of retinol in the testis leads to increased apoptosis of germ cells [42], and the testis of RAR{alpha} gene knockout mice had a similar dramatic increase in apoptosis of germ cells (unpublished results). Thus, in contrast to the critical role that RAR{alpha} plays in the testis, the very low level of RAR{alpha} found in the liver implies that it may have a minor role in the liver relative to its role in the testis. Similarly, the role of PPAR{gamma} in the liver may be less than that of PPAR{alpha} in the liver, whereas both PPAR{alpha} and PPAR{gamma} could play the major role in the testis.

When investigating the mechanisms of phthalate toxicity in the testis, the previous experimental approach has been to study the apoptosis-associated proteins, such as Fas ligand expression in Sertoli cells [35, 43], caspase-3 [44], and members of the tumor necrosis factor superfamily of death receptors, including TRAIL-R1 (DR 4) and TRAIL-R2 (DR 5) [45]. These studies clearly are important, because DEHP and MEHP increase apoptosis of germ cells. To our knowledge, however, what happens earlier, upstream of the activation of apoptosis-related proteins after PP treatment, has not been studied.

In the present study, we investigated the effect of phthalates on the activity of RAR{alpha} and PPARs, which appears to change within an hour after phthalate treatment. Previously, it was shown that nuclear localization and transcriptional activity of RAR{alpha} were coupled tightly, and both were inhibited by PPs in both primary Sertoli cells and the MSC-1 cell line, even in the presence of tRA [22]. This is significant for the function of the testis, both because RAR{alpha} is an essential receptor for testicular function and because disruption of the RAR{alpha} gene results in male sterility phenotype [23]. Conversely, PPAR{alpha}, which was distributed both in the cytoplasmic and nuclear compartments in the absence of PPs, translocated to the nucleus in the presence of MEHP or Wy 14,643 in primary rat Sertoli cells and MSC-1 cell lines [22]. This translocation was accompanied by an increase in transcriptional activity. The combination of MEHP and tRA further increased the nuclear localization of PPAR{alpha}, suggestive of tRA stimulating RXR, the permissive heterodimer partner of PPAR [22]. Similarly, in the present study, both MEHP and tRA were necessary to translocate PPAR{gamma} into the nucleus, but the addition of MEHP alone showed only a concentration of PPAR{gamma} around the nuclear envelope. Thus, this ligand requirement of the RXR partner also is the case for PPAR{gamma}. We have previously noted that this may indicate a further loss of ligand for RAR{alpha} and, thus, decreased transcriptional function for RAR{alpha} [22].

Taken together, the results suggest that if the amount of tRA is not limiting, as in Sertoli cells after 20 days of age in rats [46], both PPAR{alpha} and PPAR{gamma} could be responsive to MEHP and Wy 14,643 and translocate into the nucleus in Sertoli cells. In other words, a decrease in the amount of RAR{alpha} in the nucleus is replaced by both PPAR{alpha} and PPAR{gamma}, which are increased in the nucleus after PP treatment. Therefore, these results further support the hypothesis that PPs alter the repertoire of transcriptionally active nuclear receptors, including RAR{alpha}, PPAR{alpha}, and PPAR{gamma}, in Sertoli cells, potentially changing gene expression [22]. Moreover, these results, combined with the results concerning the relative abundance of receptors, are in line with PPs acting on the testis by mechanisms that are either dependent or independent of PPAR{alpha} [18]. Supportive of this idea, a recent study showed an increase in the expression of plasminogen activator inhibitor-1, a PPAR{gamma} target gene, in the testis compared to the liver after di-n-butyl phthalate treatment of animals [47]. However, it should be cautioned that our data do not rule out the possibility that other nonreceptor mechanisms, such as antiandrogenic [11, 18] or estrogenic mechanisms [48], are responsible for the phthalate action in the testis. It is interesting that the antiandrogenic effect of peroxisome proliferators may be mediated by PPAR{alpha} in Leydig cells [18].

Sertoli cells are not totally unique regarding the regulation of nuclear localization of PPARs. Recently, a similar dual activation of these PPARs was found in ovarian granulosa cells [49]. In addition, PPAR{alpha} was found constitutively in the cytoplasm of human macrophages, only stimulated to translocate by ligands [50], and PPAR isoforms were shown to make the cytoplasmic-to-nuclear transition on treatment with 15-deoxy{Delta} 12,14-PGJ2 in endothelial cells [51].

On the other hand, intriguingly, the cellular localization of RAR{alpha} in the liver was not susceptible to PPs, and RAR{alpha} was confined primarily to the cytoplasm. The tRA did not induce RAR{alpha} into the nucleus of ML-457 cells, and under this circumstance, MEHP had no effect on RAR{alpha} nuclear localization. Moreover, the treatment of liver cells with PPs did not elicit any changes regarding the nuclear localization of the endogenous PPAR{alpha} or PPAR{gamma}. They mainly were nuclear, with a diffuse distribution in the cytoplasm both in the presence and absence of MEHP.

Previously, it was shown that the nuclear localization of RAR{alpha} in Sertoli cells was influenced negatively by protein kinase A [32] and positively by protein kinase C and MAPK [24]. Thus, in light of a number of previous studies and their results, which demonstrated that PPs activate MAPK in liver cells [26, 27, 52], we decided to determine whether the levels of phosphorylated ERK1, ERK2, and p38 MAPK were altered after phthalate treatment of MSC-1 and primary Sertoli cells and to compare the results with those in liver cells. Similar to previous studies, we found that MAPK and p38 also were activated rather rapidly within 5–20 min in ML-457 liver cells after MEHP and Wy-14,643 treatment. This increased MAPK was demonstrated to be associated with an increased expression of immediate early genes related to growth and proliferation of liver cells [25, 26, 28, 29], but it inhibited PPAR{gamma}-mediated transcription [53, 54]. Thus, this increased MAPK activation could account for an increased proliferation of liver cells after phthalate treatment.

Remarkably, in striking contrast to liver cells, the phosphorylated MAPK levels decreased in Sertoli cells rather rapidly after PP treatments, whereas phosphorylated p38 levels were not altered. These results are compatible with MAPK activation being important for RAR{alpha} activation [24] and with PPs inhibiting the nuclear localization and transcriptional activity of RAR{alpha} within an hour in Sertoli cells [22]. However, it remains to be investigated whether deactivation of MAPK in Sertoli cells increases transcriptional function of PPAR{alpha} or PPAR{gamma} and decreases the nuclear localization and transcriptional function of RAR{alpha} in vivo, leading to impaired RAR{alpha} function in the testis, which then could account for a decreased testicular function. Moreover, it should be cautioned that the results we report likely are caused by the acute, higher-dose effects and not by the chronic, lower-dose effects. The mechanisms of phthalate action and, thus, the risk assessment may be different for an acute, higher exposure than those for a chronic, lower exposure.

In summary, we present a mechanistic model in which the initial critical event after PP treatment may be deactivation of the MAPK pathway in Sertoli cells or activation of the MAPK pathway in the liver. Coincidentally, PP exposure is linked to differential activation of RAR{alpha} and PPAR receptors in the nucleus of Sertoli cells but not with any alteration in the receptor levels in either the testis or the liver. The changes in the receptor function in the testis, but not in the liver, could cause differential gene expression in the testis and the liver that, in turn, may lead to the contrasting biological effects on testicular germ cells and liver cells. Further investigation is necessary to determine the tissue-specific changes of gene expression associated with the nuclear receptor activation or deactivation that leads to the eventual cellular effects in vivo.


   ACKNOWLEDGMENTS
 
We thank Drs. January Wahlstrom and Jeffery Jones (Department of Chemistry, Washington State University) for generously synthesizing and purifying the MEHP. We also thank Drs. Brian J. Ledwith and Cindy J. Pauley from Merck Pharmaceuticals for providing the ML-457 cell line. Finally, we thank Amy Sorg for technical assistance.


   FOOTNOTES
 
1 Supported by grant ES09978 from the National Institute of Environmental Health Sciences. Back

2 Correspondence: Kwan Hee Kim, School of Molecular Biosciences, Washington State University, Pullman, WA 99164-4234. FAX 509 335 1907; khkim{at}wsu.edu Back

Received: 3 May 2004.

First decision: 28 May 2004.

Accepted: 8 November 2004.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Kavlock R, Boekelheide K, Chapin R, Cunningham M, Faustman E, Foster P, Golub M, Henderson R, Hinberg I, Little R, Seed J, Shea K, Tabacova S, Tyl R, Williams P, Zacharewski T, NTP Center for the Evaluation of Risks to Human Reproduction: phthalates expert panel report on the reproductive and developmental toxicity of di(2-ethylhexyl) phthalate. Reprod Toxicol 2002 16:529-653[CrossRef][Medline]
  2. Kavlock R, Boekelheide K, Chapin R, Cunningham M, Faustman E, Foster P, Golub M, Henderson R, Hinberg I, Little R, Seed J, Shea K, Tabacova S, Tyl R, Williams P, Zacharewski T, NTP Center for the Evaluation of Risks to Human Reproduction: phthalates expert panel report on the reproductive and developmental toxicity of di-isononyl phthalate. Reprod Toxicol 2002 16:679-708[CrossRef][Medline]
  3. Wilkinson CF, Lamb JC IV, The potential health effects of phthalate esters in children's toys: a review and risk assessment. Regul Toxicol Pharmacol 1999 30:140-155[CrossRef][Medline]
  4. Shea KM, Pediatric exposure and potential toxicity of phthalate plasticizers. Pediatrics 2003 111:1467-1474[Abstract/Free Full Text]
  5. Albro PW, Chapin RE, Corbett JT, Schroeder J, Phelps JL, Mono-2-ethylhexyl phthalate, a metabolite of di-(2-ethylhexyl) phthalate, causally linked to testicular atrophy in rats. Toxicol Appl Pharmacol 1989 100:193-200[CrossRef][Medline]
  6. Corton JC, Lapinskas PJ, Gonzalez FJ, Central role of PPAR{alpha} in the mechanism of action of hepatocarcinogenic peroxisome proliferators. Mutat Res 2000 448:139-151[Medline]
  7. Boekelheide K, Sertoli cell toxicants. In: Russell LD, Griswold MD (eds.), The Sertoli Cell. Clearwater, FL: Cache River Press; 1993:564– 567
  8. Li LH, Heindel JJ, Sertoli cell toxicants. In: Korach KS (ed.), Reproductive and Development Toxicology. New York: Marcel Decker; 1998:655–691
  9. Li LH, Jester WF, Orth JM, Effects of relatively low levels of mono-(2-ethylhexyl) phthalate on cocultured Sertoli cells and gonocytes from neonatal rats. Toxicol Appl Pharmacol 1998 153:258-265[CrossRef][Medline]
  10. Li LH, Jester WF Jr, Laslett AL, Orth JM, A single dose of di-(2-ethylhexyl) phthalate in neonatal rats alters gonocytes, reduces Sertoli cell proliferation, and decreases cyclin D2 expression. Toxicol Appl Pharmacol 2000 166:222-229[CrossRef][Medline]
  11. Mylchreest E, Wallace DG, Cattley RC, Foster PMD, Dose-dependent alterations in androgen-regulated male reproductive development in rats exposed to di(n-butyl)phthalate during late gestation. Toxicol Sci 2000 55:143-151[Abstract/Free Full Text]
  12. Akingbemi BT, Youker RT, Sottas CM, Ge R, Katz E, Klinefelter GR, Zirkin BR, Hardy MP, Modulation of rat Leydig cell steroidogenic function by di(2-ethylhexyl)phthalate. Biol Reprod 2001 65:1252-1259[Abstract/Free Full Text]
  13. Green S, PPAR: a mediator of peroxisome proliferator action. Mutat Res 1995 333:101-109[Medline]
  14. Green S, Wahli W, Peroxisome proliferator-activated receptors: finding the orphan a home. Mol Cell Endocrinol 1994 100:149-153[CrossRef][Medline]
  15. Varanasi U, Chu R, Huang Q, Castellon R, Yeldandi AV, Reddy JK, Identification of a peroxisome proliferator-responsive element upstream of the human peroxisomal fatty acyl coenzyme A oxidase gene. J Biol Chem 1996 271:2147-2155[Abstract/Free Full Text]
  16. Gonzalez FJ, Recent update on the PPAR alpha-null mouse. Biochimie 1997 79:139-144[Medline]
  17. Ward JM, Peters JM, Perella CM, Gonzalez FJ, Receptor and nonreceptor-mediated organ-specific toxicity of di(2-ethylhexyl)phthalate (DEHP) in peroxisome proliferator-activated receptor {alpha}-null mice. Toxicol Pathol 1998 26:240-246[Medline]
  18. Gazouli M, Yao ZX, Boujrad N, Corton JC, Culty M, Papadopoulos V, Effect of peroxisome proliferators on Leydig cell peripheral-type benzodiazepine receptor gene expression, hormone-stimulated cholesterol transport, and steroidogenesis: role of the peroxisome proliferator-activator receptor {alpha}. Endocrinology 2002 143:2571-2583[Abstract/Free Full Text]
  19. Lee SS, Pineau T, Drago J, Lee EJ, Owens JW, Kroetz DL, Fernandez-Salguero PM, Westphal H, Gonzalez FJ, Targeted disruption of the {alpha} isoform of the peroxisome proliferator-activated receptor gene in mice results in abolishment of the pleiotropic effects of peroxisome proliferators. Mol Cell Biol 1995 15:3012-3022[Abstract]
  20. Peters JM, Cattley RC, Gonzalez FJ, Role of PPAR{alpha} in the mechanism of action of the nongenotoxic carcinogen and peroxisome proliferator Wy-14,643. Carcinogenesis 1997 18:2029-2033[Abstract/Free Full Text]
  21. Maloney EK, Waxman DJ, Trans-Activation of PPAR{alpha} and PPAR{gamma} by structurally diverse environmental chemicals. Toxicol Appl Pharmacol 1999 161:209-218[CrossRef][Medline]
  22. Dufour JM, Vo MN, Bhattacharya N, Okita J, Okita R, Kim KH, Peroxisome proliferators disrupt retinoic acid receptor {alpha} signaling in the testis. Biol Reprod 2003 68:1215-1224[Abstract/Free Full Text]
  23. Lufkin T, Lohnes D, Mark M, Dierich A, Gorry P, Gaub MP, LeMeur M, Chambon P, High postnatal lethality and testis degeneration in retinoic acid receptor {alpha} mutant mice. Proc Natl Acad Sci U S A 1993 90:7225-7229[Abstract/Free Full Text]
  24. Braun KW, Vo MN, Kim KH, Positive regulation of retinoic acid receptor alpha by protein kinase C and mitogen-activated protein kinase in Sertoli cells. Biol Reprod 2002 67:29-37[Abstract/Free Full Text]
  25. Ledwith BJ, Johnson TE, Wagner LK, Pauley CJ, Manam S, Galloway SM, Nichols WW, Growth regulation by peroxisome proliferators: opposing activities in early and late G1. Cancer Res 1996 56:3257-3264[Abstract/Free Full Text]
  26. Rokos CL, Ledwith BJ, Peroxisome proliferators activate extracellular signal-regulated kinases in immortalized mouse liver cells. J Biol Chem 1997 272:13452-13457[Abstract/Free Full Text]
  27. Mounho BJ, Thrall BD, The extracellular signal-regulated kinase pathway contributes to mitogenic and antiapoptotic effects of peroxisome proliferators in vitro. Toxicol Appl Pharmacol 1999 159:125-133[CrossRef][Medline]
  28. Ledwith BJ, Manam S, Troilo P, Joslyn DJ, Galloway SM, Nichols WW, Activation of immediate-early gene expression by peroxisome proliferators in vitro. Mol Carcinog 1993 8:20-27[Medline]
  29. Pauley CJ, Ledwith BJ, Kaplanski C, Peroxisome proliferators activate growth regulatory pathways largely via peroxisome proliferator-activated receptor {alpha}-independent mechanisms. Cell Signal 2002 14:351-358[CrossRef][Medline]
  30. Kaufmann W, Deckardt K, McKee RH, Butala JH, Bahnemann R, Tumor induction in mouse liver: di-isononyl phthalate acts via peroxisome proliferation. Regul Toxicol Pharmacol 2002 36:175-183[CrossRef][Medline]
  31. Favata MF, Horiuchi KY, Manos EJ, Daulerio AJ, Stradley DA, Feeser WS, Van Dyk DE, Pitts WJ, Earl RA, Hobbs F, Copeland RA, Magolda RL, Scherle PA, Trzaskos JM, Identification of a novel inhibitor of mitogen-activated protein kinase kinase. J Biol Chem 1998 273:18623-18632[Abstract/Free Full Text]
  32. Braun KW, Tribley WA, Griswold MD, Kim KH, Follicle-stimulating hormone inhibits all-trans-retinoic acid-induced retinoic acid receptor {alpha} nuclear localization and transcriptional activation in mouse Sertoli cell lines. J Biol Chem 2000 275:4145-4151[Abstract/Free Full Text]
  33. Tung PS, Skinner MK, Fritz IB, Fibronectin synthesis is a marker for peritubular cell contaminants in Sertoli cell-enriched cultures. Biol Reprod 1984 30:199-211[Abstract]
  34. Richburg JH, Boekelheide K, Mono-(2-ethylhexyl) phthalate rapidly alters both Sertoli cell vimentin filaments and germ cell apoptosis in young rat testes. Toxicol Appl Pharmacol 1996 137:42-50[CrossRef][Medline]
  35. Lee J, Richburg JH, Younkin SC, Boekelheide K, The Fas system is a key regulator of germ cell apoptosis in the testis. Endocrinology 1997 138:2081-2088[Abstract/Free Full Text]
  36. Chomczynski P, Sacchi N, Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem 1987 162:156-159[Medline]
  37. Wang Q, Fujii H, Knipp GT, Expression of PPAR and RXR isoforms in the developing rat and human term placentas. Placenta 2002 23:661-671[CrossRef][Medline]
  38. Bradford MM, A rapid and sensitive method for the quantitation of microgram quantities of protein using the principle of protein-dye binding. Anal Biochem 1976 72:248-254[CrossRef][Medline]
  39. Okita JR, Castle PJ, Okita RT, Characterization of cytochrome P450 in liver and kidney of rats treated with di-(2-ethylhexyl)phthalate. Biochem Toxicol 1993 8:135-144
  40. James NH, Soames AR, Roberts RA, Suppression of hepatocyte apoptosis and induction of DNA synthesis by the rat and mouse hepatocarcinogen diethylhexylphlathate (DEHP) and the mouse hepatocarcinogen 1,4-dichlorobenzene (DCB). Arch Toxicol 1998 72:784-790[CrossRef][Medline]
  41. Akmal KM, Dufour JM, Kim KH, Retinoic acid receptor {alpha} gene expression in the rat testis: potential role during the prophase of meiosis and in the transition from round to elongating spermatids. Biol Reprod 1997 56:549-556[Abstract]
  42. Akmal KM, Dufour JM, Vo M-N, Higginson S, Kim KH, Ligand-dependent regulation of retinoic acid receptor {alpha} in rat testis: in vivo response to depletion and repletion of vitamin A. Endocrinology 1998 139:1239-1248[Abstract/Free Full Text]
  43. Richburg JH, Nanez A, Williams LR, Embree ME, Boekelheide K, Sensitivity of testicular germ cells to toxicant-induced apoptosis in gld mice that express a nonfunctional form of Fas ligand. Endocrinology 2000 141:787-793[Abstract/Free Full Text]
  44. Dalgaard M, Nellemann C, Lam HR, Sorensen IK, Ladefoged O, The acute effects of mono(2-ethylhexyl)phthalate (MEHP) on testes of prepubertal Wistar rats. Toxicol Lett 2001 122:69-79[CrossRef][Medline]
  45. Giammona CJ, Sawhney P, Chandrasekaran Y, Richburg JH, Death receptor response in rodent testis after mono-(2-ethylhexyl) phthalate exposure. Toxicol Appl Pharmacol 2002 185:119-127[CrossRef][Medline]
  46. Cavazzini D, Galdieri M, Ottonello S, Retinoic acid synthesis in the somatic cells of rat seminiferous tubules. Biochim Biophys Acta 1996 1313:139-145[Medline]
  47. Kobayashi T, Niimi S, Kawanishi T, Fukuoka M, Hayakawa T, Changes in peroxisome proliferator-activated receptor gamma-regulated gene expression and inhibin/activin-follistatin system gene expression in rat testis after an administration of di-n-butyl phthalate. Toxicol Lett 2003 138:215-225[CrossRef][Medline]
  48. Atanassova N, McKinnell C, Turner KJ, Walker M, Fischer JS, Morley M, Millar MR, Groome NP, Sharpe RM, Comparative effects of neonatal exposure of male rats to potent and weak (environmental) estrogens on spermatogenesis at puberty and the relationship to adult testis size and fertility: evidence for stimulatory effects of low estrogen levels. Endocrinology 2000 141:3898-3907[Abstract/Free Full Text]
  49. Lovekamp-Swan T, Jetten AM, Davis BJ, Dual activation of PPAR{alpha} and PPAR{gamma} by mono-(2-ethylhexyl) phthalate in rat ovarian granulosa cells. Mol Cell Endocrinol 2003 201:133-141[CrossRef][Medline]
  50. Chinetti G, Griglio S, Antonucci M, Torra IP, Delerive P, Majd Z, Fruchart JC, Chapman J, Najib J, Staels B, Activation of proliferator-activated receptors {alpha} and {gamma} induces apoptosis of human monocyte-derived macrophages. J Biol Chem 1998 273:25573-25580[Abstract/Free Full Text]
  51. Bishop-Bailey D, Hla T, Endothelial cell apoptosis induced by the peroxisome proliferator-activated receptor (PPAR) ligand 15-deoxy-12, 14-prostaglandin J2. J Biol Chem 1999 274:17042-17048[Abstract/Free Full Text]
  52. Passilly P, Schohn H, Jannin B, Malki MC, Boscoboinik D, Dauca M, Latruffe N, Phosphorylation of peroxisome proliferator-activated receptor {alpha} in rat Fao cells and stimulation by ciprofibrate. Biochem Pharmacol 1999 58:1001-1008[CrossRef][Medline]
  53. Hu E, Kim JB, Sarraf P, Spiegelman BM, Inhibition of adipogenesis through MAP kinase-mediated phosphorylation of PPAR{gamma}. Science 1996 274:2100-2103[Abstract/Free Full Text]
  54. Chan GK, Deckelbaum RA, Bolivar I, Goltzman D, Karaplis AC, PTHrP inhibits adiopocyte differentiation by down-regulating PPAR{gamma} activity via a MAPK-dependent pathway. Endocrinology 2001 142:4900-4909[Abstract/Free Full Text]