Peroxisome Proliferator–Activated Receptor α Protects against Glomerulonephritis Induced by Long-Term Exposure to the Plasticizer Di-(2-Ethylhexyl)Phthalate

Animals and DEHP Treatment

PPARα-null and wild-type mice were on a SV/129 genetic background, as described elsewhere (16). These mice were maintained in a facility that was free of specific pathogens according to Shinshu University and National Institutes of Health animal care guidelines and Accreditation of Laboratory Animal Care guidelines. The mice were housed in a temperature- and light-controlled environment (25°C; 12-h light/dark cycle) and maintained on stock rodent diet and tap water ad libitum until reaching 12 wk of age. Twelve-week-old male wild-type and PPARα-null mice (body weight for both 25 to 30 g) were fed their regular diet or a DEHP-containing diet (0.01 or 0.05%) for 22 mo. The beginning and ending sizes of each group were as follows: n = 25 and 24 (one mouse died) for the control wild-type group; n = 25 and 23 (two mice died) for the 0.01% DEHP wild-type group; n = 21 and 20 (one mouse died) for the 0.05% DEHP wild-type group; n = 26 and 25 (one mouse died) for the control PPARα-null group; n = 28 and 25 (three mice died) for the 0.01% DEHP PPARα-null group; and n = 34 and 31 (three mice died) for the 0.05% DEHP PPARα-null group. The clinical parameters of each group of mice were checked at 0, 6, 12, and 22 mo during the experimental period. Systolic BP was measured by a programmed sphygmomanometer (BP-98A; Softron Corp., Tokyo, Japan) using a tail-cuff method. Urine protein was measured as described previously (10). Serum urea nitrogen and creatinine were determined by a clinical analyzer (JCA-BM2250; JEOL, Tokyo, Japan). All of the mice that had survived the 22-mo of experimental period were killed after finishing DEHP treatment. One PPARα-null mouse that was exposed to 0.05% DEHP exhibited marked hydronephrosis and therefore was not used in the histopathologic and biochemical analyses. The other mice showed no obvious abnormal macroscopic features, and the kidneys from these mice were used in the following analyses.

Histopathologic Analyses

Tissues from kidneys in each group of mice were fixed in 4% paraformaldehyde; embedded in paraffin; sectioned; and stained with hematoxylin and eosin, periodic acid-Schiff, or periodic acid-methenamine-silver for histopathologic examination using light microscopy. For semiquantitative histologic analyses, more than 20 glomeruli from each kidney section were examined. Degrees of cell proliferation and mesangial expansion were estimated using a scale that ranged from 0 to 3 (0, normal; 1, mild; 2, moderate; 3, severe). Indices were calculated using the following formula: Index = (n₀× 0) + (n₁× 1) + (n₂× 2) + (n₃× 3)/∑_n (∑_n > 20). These histopathologic analyses were performed in a blinded manner by two observers who were unaware of the study protocol. Cryosections for immunofluorescent analyses were stained using FITC-labeled anti-mouse C3, IgG, IgA, or IgM antibodies (ICN Pharmaceuticals, Aurora, OH). Paraffin sections were deparaffinized and stained using an indirect immunoperoxidase technique. A primary mAb to a mouse macrophage marker, F4/80 antigen, was purchased from BMA Biomedicals (Augst, Switzerland). Polyclonal primary antibodies to two oxidative stress markers, 4-hydroxynonenal (4-HNE) and 8-hydroxy-2′-deoxyguanosine (8-OHdG), respectively, were purchased from Alexis Corp. (Lausen, Switzerland) and Chemicon International (Temecula, CA). Tissues to be used for electron microscopy were fixed immediately in 2.5% glutaraldehyde, osmicated, dehydrated in increasing graded ethanol concentrations, and embedded in Epon resin. Ultrathin sections were double stained with uranyl acetate and lead citrate and examined with a JEM 1200EX II electron microscope (JEOL, Tokyo, Japan).

Immunoblot Analyses

For immunoblot analyses, glomeruli were isolated from all kidneys in each group of mice, using a previously described sieving method (17). Glomerular extracts were subjected to 9 to 15% SDS-PAGE and then transferred to nitrocellulose membranes. These membranes were incubated with primary antibody, followed by incubation with alkaline phosphatase–conjugated secondary antibody. Polyclonal primary antibodies to catalase were prepared using purified catalase (18). Primary antibodies to α-smooth muscle actin (α-SMA) and 4-HNE were purchased from DakoCytomation (Glostrup, Denmark) and Alexis Corp, respectively. Other primary antibodies, against proliferating cell nuclear antigen, TGFβ1, Nox4, p47phox, Cu,Zn-superoxide dismutase (SOD), Mn-SOD, and glutathione peroxidase, were obtained from Santa Cruz Biotechnology (Santa Cruz, CA).

Analyses of mRNA

Analyses of mRNA were performed using a real-time PCR. One microgram of total RNA was extracted from the isolated glomeruli of each group and reverse-transcribed using oligo(dT) primers and Superscript reverse transcriptase (Invitrogen, Carlsbad, CA). The cDNA were quantified with an ABI PRISM 7700 sequence detection system (Applied Biosystems, Foster City, CA), using specific primers and SYBR Green double-stranded DNA (dsDNA) binding dye I. Specific primers were designed as follows: 5′-CCTCAGGGTACCACTACGGAGT-3′ and 5′-GCCGAATAGTTCGCCGAA-3′ for PPARα (GenBank accession no. NM_011144); 5′-TTCCACTATGGAGTTCATGCTTGT-3′ and 5′-TCCGGCAGTTAAGATCACACCTA-3′ for PPARγ (NM_011146); 5′-CACCTGCAAGACCATCGACAT-3′ and 5′-TGGCGAGCCTTAGTTTGGA-3′ for TGFβ1 (NM_011577); 5′-GCCCCGCACAGCCATGTTTCAG-3′ and 5′-CATGGAGTCCAGGCCGCTGTCGTG-3′ for IκBα (U36277); 5′-TGACCCCCAAGGCTCAAATATG-3′ and 5′-ACCCAGGTCCTCGCTTATGAT-3′ for cyclooxygenase2 (NM_011198); 5′-TCCGGACTTTCGATCTTCCA-3′ and 5′-GAGCTTCAGAGGCAGGAAACA-3′ for intercellular adhesion molecule 1 (M31585), 5′-CAGCCGATGGGTTGTACCTT-3′ and 5′-GTGGGTGAGGAGCACGTAGTC-3′ for TNFα (NM_013693); 5′-CGTCCTGACAATGCAGACCTT-3′ and 5′-CCCCATGAAACGCATGAACT-3′ for TNF receptor 1 (TNR1) (M60468). Glyceraldehyde-3-phosphate dehydrogenase was used as the internal control for PCR amplification.

Measurement of Serum Mono(2-Ethylhexyl)Phthalate Concentration

Serum mono(2-ethylhexyl)phthalate (MEHP) concentration was measured as described previously (19). Briefly, MEHP was extracted from serum samples with ethyl acetate. N-Methyl-N-(tert-butyl-dimethylsilyl)trifluoroacetamide (GL Sciences, Tokyo, Japan) was added to the MEHP extracts and left at room temperature for 60 min. The MEHP tert-buthyl-dimethylsilyl derivative was analyzed by gas chromatography with mass-selective detection (6890 N, 5973 N; Agilent Technologies, Santa Clara, CA). Serum samples of hemodialysis (HD) patients and healthy volunteers were collected in July 2006 with written informed consent. This study was conducted in accordance with the Declaration of Helsinki. Signed informed consent to participate in the study was obtained from all of the patients, and the study protocols were approved by the Medical Ethics Committee of the Shinshu University School of Medicine.

Measurements of Anti-dsDNA Antibody and 50% Hemolytic Complement Activity

The titer of serum anti-dsDNA antibody was determined by ELISA as shown previously (20). The standard 50% hemolytic complement activity (CH50) was measured as described previously (21).

Statistical Analyses

Analysis of significant differences with respect to interactive effects of two factors (PPARα gene status and DEHP treatment) was performed using a two-way ANOVA. P < 0.05 was used as the measure of significance.

DEHP Exposure Levels and Serum MEHP Concentrations

Food consumption during the experimental period was uninterrupted and similar in all groups (3.1 ± 0.7 g/d per mouse). Mean daily ingestion of DEHP approximated the maximal medical exposure in humans (8 to 11 mg/kg body wt per d for the 0.01% DEHP groups of wild-type and PPARα-null mice; 42 to 55 mg/kg body wt per d for the 0.05% DEHP groups of the two genotypes; <22.6 mg/kg body wt per d for patients undergoing therapy using PVC-containing medical devices [2]). Because most ingested DEHP is hydrolyzed efficiently by lipases to MEHP, a major DEHP metabolite whose toxicity is much more intensive than that of DEHP (2), the serum MEHP concentration in each group of mice at 22 mo was measured. The concentration increased in a DEHP dosage-dependent manner in DEHP-exposed mice of both genotypes, and there was no difference between the genotypes statistically (0.047 ± 0.014 and 0.041 ± 0.007 μg/ml for the control groups of wild-type [n = 24] and PPARα-null mice [n = 25], respectively; 0.370 ± 0.097 and 0.429 ± 0.140 μg/ml for the 0.01% DEHP groups of wild-type [n = 23] and PPARα-null mice [n = 25], respectively; 1.404 ± 0.371 and 1.737 ± 0.689 μg/ml for the 0.05% DEHP groups of wild-type [n = 20] and PPARα-null mice [n = 30], respectively). These findings suggest that the DEHP exposure levels that were chosen in this study well influence the serum concentrations of the major active metabolite, MEHP. Serum concentrations of MEHP in HD patients, who are exposed frequently to DEHP through HD sessions, were unexpectedly lower (0.056 ± 0.018 μg/ml [0.016 to 0.110 μg/ml] for the HD patients after the HD sessions [n = 109]; 0.014 ± 0.003 μg/ml for the healthy volunteers [n = 16]) than those in DEHP-exposed mice. Therefore, these findings indicate that serum MEHP concentrations probably are influenced by a species difference in DEHP metabolic efficiency.

Systemic Effects of Long-Term Dietary Exposure to DEHP

The systemic effects in all groups of mice were followed up throughout the experimental period. The levels of systolic BP, daily urinary protein excretion, serum urea nitrogen, and serum creatinine did not differ among the groups until 6 mo. Systolic BP and daily urinary protein excretion in DEHP-exposed PPARα-null mice increased in time-dependent and DEHP dosage-dependent manners after 6 mo (Figure 1, A and B). Outstanding increases of urine protein excretion in 0.01 and 0.05% DEHP-exposed PPARα-null mice were observed at both 12 and 22 mo, but the development of hypertension in 0.01% DEHP-exposed PPARα-null mice was obscure (Figure 1, A and B). Therefore, hypertension may have begun to develop after proteinuria. At 22 mo, obvious increases in serum urea nitrogen and serum creatinine in DEHP-exposed PPARα-null mice were observed in a DEHP dosage-dependent manner (Figure 1, C and D). Systolic BP and daily urinary protein excretion in exposed wild-type mice slightly increased at 22 mo, but serum urea nitrogen and serum creatinine did not change at any time during the experimental period. These data suggest DEHP-dependent renal dysfunction especially in exposed PPARα-null mice.

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Figure 1.

Effects of a diet that included di-(2-ethylhexyl)phthalate (DEHP; 0.01 or 0.05%) during 22 mo in wild-type (WT) and peroxisome proliferator–activated receptor α (PPARα)-null (KO) mice. Systolic BP (A), daily urinary protein excretion (B), serum urea nitrogen (C), and serum creatinine (D) were measured. •, control WT group (n = 24); ▪, 0.01% DEHP WT group (n = 23); ▴, 0.05% DEHP WT group (n = 20); ○, control KO group (n = 25); □, 0.01% DEHP KO group (n = 25); ▵, 0.05% DEHP KO group (n = 30). Data are means ± SD. *P < 0.05, **P < 0.01, ***P < 0.001, significantly different from the respective control group; ^#P < 0.05, ^##P < 0.01, ^###P < 0.001, significant differences between WT and KO mice.

In all groups, body weight, kidney weight, and testicular weight were not different among the groups; however, liver weight in exposed wild-type mice at 22 mo decreased in a dosage-dependent manner (liver weight/body weight 4.28 ± 0.64% for the control wild-type group, 3.98 ± 0.45% for the 0.01% DEHP wild-type group, and 3.92 ± 0.26% for the 0.05% DEHP wild-type group). Because this effect was not observed in PPARα-null mice, the decrease in liver weight most likely reflected PPARα-dependent long-term hepatotoxicity. Because symptoms that were related to renal dysfunction were prominent, we focused on renal toxicity in the subsequent analyses.

DEHP Induces Immune-Complex Glomerulonephritis

The cause of renal dysfunction that was observed in these mice was examined by histopathologic analyses. Light microscopy using periodic acid-methenamine-silver staining demonstrated outstanding glomerular lesions with cellular proliferation and mesangial expansion in all of DEHP-exposed PPARα-null mice at 22 mo (Figure 2A). Cell proliferation and mesangial expansion in exposed PPARα-null mice increased in a dosage-dependent manner (Figure 2, B and C). Approximately 25% of PPARα-null mice that were exposed to 0.05% DEHP showed severe inflammatory findings such as mesangiolysis, mesangial edema, crescent formation, and macrophage infiltration (Figure 2, D and E). In contrast, only mild glomerular lesions were seen in DEHP-exposed wild-type mice. These findings indicate that long-term exposure to DEHP induces DEHP-dependent glomerulonephritis in PPARα-null mice.

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Figure 2.

Light microscopic analyses of glomerular lesions. (A) Representative glomeruli from DEHP-exposed WT and KO mice at 22 mo of experiment. Sections were stained with periodic acid-methenamine-silver (PAM). Bar = 20 μm. (B and C) Indices of glomerular lesions (cell proliferation index and mesangial expansion index) at 22 mo. Data are means ± SD (n = 24 for the control WT group; n = 23 for the 0.01% DEHP WT group; n = 20 for the 0.05% DEHP WT group; n = 25 for the control KO group; n = 25 for the 0.01% DEHP KO group; n = 30 for the 0.05% DEHP KO group). **P < 0.01, ***P < 0.001, significantly different from the respective control group; ^##P < 0.01, ^###P < 0.001, significant differences between WT (▪) and KO (□) mice. (D) Mesangiolytic and hypercellular inflammatory findings representing the glomeruli in 25% of KO mice that were exposed to 0.05% DEHP. Bar = 20 μm. (E) Immunohistochemical analysis of glomeruli in a KO mouse that was exposed to 0.05% DEHP. Deparaffinized sections were stained by an indirect immunoperoxidase technique using antibodies against mouse F4/80 (a mouse macrophage marker). Macrophages are present in intraglomerular and extracapillary spaces. Bar = 20 μm.

Light microscopy also demonstrated focal tubulointerstitial lesions with tubular atrophy and inflammatory cell infiltration in DEHP-exposed PPARα-null mice. These tubulointerstitial lesions were localized around the sclerotic glomerular lesions (Figure 3). The lesions, reported in earlier studies, such as cystic tubular dilation, tubular necrosis, tubular pigmentation, and papillary mineralization (14,22), were scarcely detected here. These findings suggest that secondary tubulointerstitial lesions developed after glomerular damages occurred.

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Figure 3.

Light microscopic analyses of tubulointerstitial lesions. Representative tubulointerstitial findings from DEHP-exposed WT and KO mice at 22 mo of experiment. Sections were stained with PAM. Bar = 100 μm.

For further characterization of DEHP-induced glomerulonephritis in PPARα-null mice, immunofluorescent and electron microscopic analyses were performed. Immunofluorescence staining demonstrated C3, IgG, IgA, and IgM deposits along peripheral glomerular capillaries in DEHP-exposed PPARα-null mice at 22 mo (Figure 4A). Electron microscopic analysis revealed massive subepithelial electron-dense deposits and diffuse foot process effacement in DEHP-exposed PPARα-null mice (Figure 4B). A small number of mesangial and subendothelial deposits also were detected. In age-matched unexposed PPARα-null mice, immunofluorescence staining indicated only a few deposits in glomeruli, and electron microscopic analysis showed no abnormal changes in podocytes. These findings suggest that this glomerulonephritis involves immune mechanisms that are affected by DEHP treatment. In each mouse group, there was no appearance of anti-dsDNA antibody and no decrease in CH50. Therefore, DEHP’s affecting glomerular immune-complex deposition seemed to act not systemically but renal specifically.

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Figure 4.

Characterization of DEHP-induced glomerulonephritis. (A) Immunofluorescence microscopic analyses of glomeruli in control and 0.05% DEHP-exposed KO mice at 22 mo of experiment. Cryosections were stained using FITC-labeled anti-mouse C3, IgG, IgA, or IgM antibodies. Bar = 20 μm. (B) Electron microscopic analysis of glomeruli in control and 0.05% DEHP-exposed KO mice at 22 mo. Arrows indicate massive subepithelial electron-dense deposits. Black arrowheads indicate mesangial deposits. White arrowhead indicates subendothelial deposits. Podocytic cytoplasm in exposed KO mice shows irregular foot process effacement. Me, mesangial cell. Bars = 2 μm. (C) Immunoblot analyses of α-smooth muscle actin (α-SMA), proliferating cell nuclear antigen (PCNA), and TGFβ1. One hundred micrograms of glomerular lysate protein that was obtained from all kidneys in each group of mice at 22 mo was used. Blots and densitometry were performed in triplicate. Data are means ± SD. *P < 0.05, **P < 0.01, ***P < 0.001, significantly different from the respective control group; ^##P < 0.01, ^###P < 0.001, significant differences between WT (▪) and KO (□) mice.

For further characterization of the glomerular lesions, known mediators of mesangial cell proliferation and fibrosis, specifically, α-SMA, proliferating cell nuclear antigen, and TGFβ1, were measured by immunoblotting. Expression of these proteins was increased markedly in PPARα-null mice at 22 mo in a DEHP dosage-dependent manner but increased only slightly in wild-type mice (Figure 4C). These findings are in agreement with the pathologic findings.

DEHP Elevates Oxidative Stress in Glomeruli

Because some studies have suggested that DEHP-induced elevation of oxidative stress contributes to hepatotoxicity and testicular toxicity (23–25), the influence of oxidative stress in DEHP-induced glomerulonephritis was examined. Immunoblot analysis showed that total amounts of 4-HNE–modified proteins, a lipid peroxidation marker, were increased markedly and dosage dependently in glomeruli of DEHP-exposed PPARα-null mice at 22 mo but increased only mildly in exposed wild-type mice (Figure 5A). By immunohistochemical analyses, the 4-HNE–modified proteins were detected in large numbers of podocytes in the glomeruli of DEHP-exposed PPARα-null mice at 22 mo (Figure 5B). These podocytes also contained 8-OHdG, an oxidative DNA damage marker (Figure 5C). These oxidative stress markers also showed slight positivity in parietal epithelial cells and proximal tubules of DEHP-exposed PPARα-null mice. Next, glomerular protein expression levels of two superoxide-generating NADPH oxidase subunits, Nox4 and p47phox, were examined. These proteins were increased markedly and dosage dependently in DEHP-exposed PPARα-null mice at 22 mo; these increases were present but reduced considerably in wild-type mice (Figure 6A). The findings suggest an increase in the generation of reactive oxygen species (ROS) in glomeruli. To this end, glomerular antioxidant protein expression was measured in each group at 22 mo. Constitutive expression of catalase, Cu,Zn-SOD, and Mn-SOD was lower in unexposed PPARα-null mice than in wild-type mice; DEHP exposure further reduced glomerular expression of these proteins in PPARα-null mice (Figure 6B). Glutathione peroxidase-1 expression remained constant in all groups. These findings suggest that DEHP exposure increases oxidative stress in glomeruli, especially in podocytes, and this effect is intensified greatly in PPARα-null mice, reflecting their intense NADPH oxidase response and low antioxidant capabilities.

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Figure 5.

Changes in glomerular amounts of oxidative stress markers in DEHP-exposed WT and KO mice. (A) Immunoblot analysis of 4-HNE–modified proteins. Twenty micrograms of glomerular lysate protein that was obtained from all kidneys in each group of mice at 22 mo of experiment was used. Blotting and densitometry were performed in triplicate. Data are means ± SD. ***P < 0.001, significantly different from the respective control group; ^###P < 0.001, significant difference between WT (▪) and KO (□) mice. (B and C) Immunohistochemical analyses of glomeruli in control and 0.05% DEHP-exposed KO mice at 22 mo. Deparaffinized sections were stained by an indirect immunoperoxidase technique using anti–4-HNE or anti–8-OHdG antibodies. Bars = 20 μm.

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Figure 6.

Changes in amounts of glomerular oxidative stress–related factors in DEHP-exposed WT and KO mice. (A and B) Immunoblot analyses of Nox4, p47phox, catalase, Cu,Zn-superoxide dismutase (Cu,Zn-SOD), Mn-SOD, and glutathione peroxidase (GPx-1). Twenty micrograms of glomerular lysate protein that was obtained from all kidneys in each group of mice at 22 mo of experiment was used. Blotting and densitometry were performed in triplicate. Data are means ± SD. **P < 0.01, ***P < 0.001, significantly different from the respective control group; ^##P < 0.01, ^###P < 0.001, significant differences between WT (▪) and KO (□) mice.

PPARα Exerts Antinephritic Effects

For investigation of the mechanism underlying development of glomerulonephritis in DEHP-exposed PPARα-null mice, expressions of mRNA encoding mediators of inflammation and fibrosis were examined, using glomerular samples at 22 mo. Because PPARα and PPARγ show anti-inflammatory effects in many tissues (13), the mRNA that encodes these transcription factors were measured using real-time PCR. As expected, glomerular expression of PPARα in PPARα-null mice remained almost undetectable throughout the experiment. Conversely, constitutive glomerular expression of PPARα in wild-type mice was high and increased in a DEHP dosage-dependent manner (Figure 7A). Constitutive glomerular expression of mRNA that encodes PPARγ, which was much less than that of PPARα in wild-type mice, did not differ between the genotypes and was decreased slightly by DEHP exposure in mice of both genotypes (Figure 7B). These findings suggest that PPARγ is not important in development of DEHP-induced glomerulonephritis. Because it was reported recently that hepatic expression of fibrogenic growth factor mRNA was influenced by PPARα expression (26), TGFβ1 mRNA was measured. Expression increased markedly in DEHP-exposed PPARα-null mice in a dosage-dependent manner, whereas the increases were only marginal in wild-type mice (Figure 7C). This mRNA expression pattern was consistent with the changes that were observed at the protein level (Figure 4C). Because PPARα is known to exert anti-inflammatory effects by inducing expression of IκBα, which antagonizes NFκB signaling (27), the expression of IκBα was measured. This mRNA was found to remain constant in exposed PPARα-null mice but increased dosage dependently in exposed wild-type mice (Figure 7D). These results were compatible with the observed pattern of PPARα mRNA expression. Next, the expression of mRNA that encode three proinflammatory mediators—cyclooxygenase 2, intercellular adhesion molecule 1, and TNFα, known as target molecules of the NFκB signaling pathway—were determined. Expression of the proinflammatory genes was increased considerably and dosage dependently in DEHP-exposed PPARα-null mice but was increased only mildly in exposed wild-type mice (Figure 7, E through G). In addition, the expression of TNR1 mRNA was measured to evaluate the sensitivity to the inflammatory response in PPARα-null mice. Surprising, constitutive expression of TNR1 was markedly higher in PPARα-null mice than in wild-type mice; this enhanced expression remained constant irrespective of exposure to DEHP (Figure 7H). These results suggest that PPARα exerts antinephritic effects by antagonizing the NFκB signaling pathway, preventing fibrosis, and lowering inflammatory sensitivity.

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Figure 7.

Changes in glomerular mRNA expression in DEHP-exposed WT and KO mice. (A through H) Analyses of mRNA in isolated glomeruli that were obtained from all kidneys in each group of mice at 22 mo of experiment. PPARα (A), PPARγ (B), TGFβ1 (C), IκBα (D), cyclooxygenase 2 (COX2; E), intercellular adhesion molecule 1 (ICAM1) (F), TNFα (G), and TNF receptor 1 (TNR1; H). The mRNA were quantified by real-time PCR. Glyceraldehyde-3-phosphate dehydrogenase mRNA was used as an internal control. Data are means ± SD of triplicate experiments. *P < 0.05, **P < 0.01, ***P < 0.001, significantly different from the respective control group; ^#P < 0.05, ^##P < 0.01, ^###P < 0.001, significant differences between WT (▪) and KO (□) mice.