Abstract
ABSTRACT. In nephrotic syndrome, large quantities of albumin enter the kidney tubule. This albumin carries with it a heavy load of fatty acids to which the proximal tubule cells are exposed at high concentration. It is postulated that exposure to fatty acids in this way is injurious to proximal tubule cells. This study has examined the ability of fatty acids to interact with peroxisome proliferator–activated receptors (PPAR) in primary cultures of human proximal tubule cells. Luciferase reporter assays in transiently transfected human proximal tubule cells were used to show that albumin bound fatty acids and other agonists activate PPARγ in a dose-dependent manner. One of the consequences of this activation is apoptosis of the cells as determined by changes in cell morphology, evidence of PARP cleavage, and appearance of DNA laddering. Overexpression of PPARγ in these cells also results in enhanced apoptosis. Both fatty acid–induced PPAR activation and apoptosis in these cells can be blocked by PPAR response element decoy oligonucleotides. Activation of PPARγ by the specific agonist PGJ2 is associated with inhibition of cell proliferation, whereas activation by albumin bound fatty acids is accompanied by increased proliferation. However, the net balance of apoptosis/proliferation favors deletion of cells. These results implicate albumin-bound fatty acids as important mediators of tubular injury in nephrosis and provide fresh impetus for pursuit of lipid-lowering strategies in proteinuric renal disease. E-mail: njb18@le.ac.uk
In patients with renal disease, the presence of proteinuria is an adverse prognostic sign such that those individuals with proteinuria are more likely to progress to end-stage renal failure than those with absent or very modest proteinuria (1,2). Histopathologic observations reveal a very close correlation between progressive renal functional impairment and tubulointerstitial inflammation, scarring, and fibrosis (3). The most prevalent protein in the urine of nephrotic patients is albumin, and several authors have postulated that albumin may have a detrimental effect on tubulointerstitial function when filtered in excess into the proximal tubule. Thus in vivo proteinuria has been associated with proliferation and apoptosis of proximal tubule cells (PTC) and interstitial inflammation (4,5). In vitro albumin stimulates various intracellular signaling pathways in PTC and induces them to produce various chemoattractants (6–10). Some authors have demonstrated a pro-apoptotic effect of albumin in cultured PTC, and others demonstrate an opposite effect (11,12).
Although there now seems little doubt that albumin can alter the properties of PTC in a number of ways, whether albumin per se is a major mediator of renal tubulointerstitial disease remains controversial. One common criticism leveled at the albumin-mediated renal damage hypothesis is the example of minimal change disease. Individuals with this condition manifest heavy proteinuria, yet, in contrast to those with other nephrotic states, they generally fail to develop tubulointerstitial disease and renal impairment. Urinary albumin in minimal change disease displays a crucial difference to that found in other nephroses being relatively devoid of fatty acids (13). Thus it is likely that albumin-bound fatty acids (FA) may be important mediators of renal tubulointerstitial disease. Although large-scale trials of lipid-lowering strategies on the progression of renal impairment in renal disease have not been performed, a recent meta-analysis of several smaller trials suggests a renoprotective effect of lipid reduction equivalent to that observed for angiotensin-converting enzyme (ACE) inhibitors in proteinuric conditions (14).
Circulating albumin is able to bind and act as a carrier for many serum-derived molecules. In particular, albumin possesses five to seven high-affinity binding sites for FA, and these FA are carried with albumin into the proximal tubule (15,16). Long-chain FA are very poorly soluble, and filtered albumin thus has the capacity to present these molecules to PTC in an unregulated manner at concentrations far above that normally determined by their solubility in aqueous solution. Furthermore, in nephrosis the FA load per albumin molecule is markedly increased, with a molar ratio of approximately 6 (FA:albumin) compared with a ratio of ≤1 in health (17). Lipid accumulation in PTC can be demonstrated by Oil Red O staining (18). This lipid accumulation results in considerable metabolic perturbation (18). Proximal tubular segments exposed to FA-bearing albumin but not FA-free albumin elaborate a lipid chemoattractant that has a potentially key role in the induction of interstitial inflammation (19).
The peroxisome proliferator activated receptors (PPAR) are a family of ligand activated transcription factors belonging to the nuclear receptor superfamily (20,21). Three subtypes, PPARα, PPARβ (also known as PPARδ or Nuc-1), and PPARγ, are found in mammalian cells, and all subtypes have been identified in the kidneys of humans and other species (20,22–26). Upon ligand binding, PPAR form heterodimers with one of the three retinoid X receptor proteins, which then bind to PPAR response elements (PPRE) within the promoter regions of target genes (27).
PPARα is highly expressed in tissues such as liver, muscle, heart, kidney, and brown adipose tissue that possess high catabolic rates for FA (20,21,28). PPARβ is ubiquitously expressed, but its functions are less well defined (20,21). PPARγ is also widely expressed, but at particularly high levels in adipose tissue, where it is a critical regulator of adipocyte differentiation (20,21,29). In addition, PPARγ agonists have been shown to induce apoptosis in fibroblasts, macrophages, and endothelial cells (30–33).
A variety of endogenous and exogenous ligands for PPAR have been identified (34). In particular, PPARα is the target for the hypolipidemic fibrate drugs, and PPARγ is the target for the antidiabetic thiazolidinediones. With a few exceptions, most FA can also activate PPAR. A number of eicosanoids have also been shown to activate PPAR, e.g., 5,8,11,14 eicosatetraynoic acid (ETYA) for PPARα and 15-deoxy-Δ(12,14)-prostaglandin J2 for PPARγ.
The functions of PPAR in the kidney have not been well studied. In view of our interest in albumin bound lipids in the pathogenesis of renal disease we have examined the potential for such FA to stimulate PPAR activity in human PTC (HPTC). We have also studied the functional consequences of these interactions, particularly as related to PTC growth and survival that are known to be abnormal in proteinuric states.
Materials and Methods
Materials
Fugene 6 transfection reagent was obtained from Roche Diagnostics (Lewes, England). The PPAR agonists ETYA and 15-deoxy-Δ(12,14)-prostaglandin J2 (PGJ2) were from Affiniti Research Products (Exeter, England). All primary anti-sera were from Santa Cruz Biotechnology Inc. (Santa Cruz, CA) and supplied by Insight Biotechnology (London, England). The plasmids, pCMX-PPARγ encoding mouse PPARγ1 and pCMX-RXRα encoding mouse retinoic acid X receptor α, were kindly provided by Dr R. Evans (Salk Institute, San Diego, CA). The plasmid pSG5-PPARα was kindly provided by Dr F. Gonzalez (National Cancer Institute, Bethesda, MD), and plasmid pSG5-PPARδ was kindly provided by Dr P. Grimaldi (Center de Biochmie, Nice, France). The reporter plasmid pPPRE-TK-luc was kindly provided by Dr M. Lazar (University of Pennsylvania, Philadelphia, PA). The green fluorescence protein encoding vector pEGFP-C1 was obtained from Clontech (Palo Alto, CA). Oligonucleotides were synthesized by Life Technologies BRL (Paisley, Scotland). Luciferase assays were performed using the LucLite kit (Packard, Pangbourne, England). β-Galactosidase assay kits were obtained from Promega (Madison, WI). Human serum albumin fraction V powder, replete with FA, was obtained from Sigma (catalogue number A1653). Essentially FA-free human serum albumin, prepared from A1653, was also obtained from Sigma (catalogue number A1887).
Cell Culture
Primary cultures of HPTC were used in all experiments. These cells were isolated as previously reported (35), with a modification of the method described by Detrisac et al. (36). In brief, the outer cortex was dissected from the normal pole of kidneys removed for the treatment of renal carcinoma. The fibrous capsule was removed, and tissue from the cortex was cut into small pieces and digested in type II collagenase (1.0 mg/ml) at 37°C for 30 min. After digestion, the cell suspension was passed through a series of sieves of diminishing mesh size. Tubular fragments passing through the sieves were seeded into 75-cm2 flasks (Costar, UK) that had been coated with bovine collagen type I and FCS proteins. The cells were grown in serum-free DMEM:F12 (Life Technologies, UK) supplemented by the addition of 25 mM HEPES buffer, 5 μg/ml insulin, 5 μg/ml transferrin, 5 ng/ml selenium, 4 pg/ml triiodo-thyronine, 36 ng/ml hydrocortisone, 100 IU/ml benzyl penicillin, and 50 μg/ml streptomycin. The cells reached confluence in 10 to 14 d and were then subcultured. Cultured cells were characterized as HPTC as reported previously (35). HPTC were never used from frozen stocks and were subcultured only up to passage 4 and then discarded.
Transient Transfection of HPTC
All transient transfections were performed in HPTC at approximately 50% confluence using Fugene 6 transfection reagent according to the manufacturer’s instructions. In preliminary experiments, cells growing on glass coverslips were transiently transfected with pEGFP-C1, fixed in paraformaldehyde at various times after transfection, and examined by fluorescence microscopy to ascertain levels of transfection efficiency.
For luciferase reporter experiments, cells were grown in non–collagen-coated 6-, 24-, or 96-well plates. HPTC were transfected with pPPRE-TK-luc and various combinations of pSVβgal, pCMX-RXRα, and pSG5-PPARα pSG5-PPARδ or pCMX-PPARγ. For 24-well plates, 0.25 μg of each plasmid DNA was used per well, and the quantity of Fugene 6 used was varied according to the quantity of plasmid DNA exactly as in the manufacturer’s instructions. For larger- and smaller-well plates, the quantity of plasmid DNA used in the transfections was increased or decreased as appropriate. In experiments involving the transfection of multiple plasmids, equivalent concentrations of DNA were maintained by the addition of appropriate empty plasmid vector.
Plasmid DNA/Fugene 6 mixtures were added to cells growing in fully supplemented media. After 24 h, media was aspirated from cells and replaced with DMEM:F12 growth media lacking insulin, transferrin, selenite tri-iodothyronine, and hydrocortisone (quiescence media) but containing various concentrations of PPAR agonists. After a additional 24 h, this media was removed and cells were lysed in a buffer consisting of 500 mM HEPES, 2% Triton N101, 1 mM CaCl2, 1 mM MgCl2, pH 7.8. Cell lysis was allowed to proceed for 10 min, a 50 μl aliquot of lysate was removed for β-galactosidase assay, and the remainder was used for luciferase assay using LucLite assay kit. Luciferase activity was measured directly in plate wells using a LumiCount luminometer (Packard, Pangbourne, England). Luciferase activity in all experiments was normalized to β-galactosidase content.
To confirm overexpression of PPAR in transiently transfected HPTC cells were lysed in SDS lysis buffer. Lysates were subjected to polyacrylamide gel electrophoresis, and separated proteins were transferred to nitrocellulose membrane. Membranes were then probed with polyclonal anti-PPARα, anti-PPARδ, or anti-PPARγ antisera, and bound primary antibody were detected by enhanced chemiluminescence (ECL, Amersham, England).
Assessment of Cell Morphology and Apoptosis
Morphology of HPTC growing in 12-well plates was examined using a Zeiss Axiovert 10 inverted fluorescence microscope. Unstained cells were studied using differential interference contrast (Normaski) microscopy. Inverted fluorescence microscopy of undisturbed HPTC cultures, to which acridine orange dye at a final concentration of 10 μg/ml had been added, was used to quantify apoptosis. Apoptotic cells were readily identifiable by shrinkage and condensation of chromatin. This technique has previously been validated and used to detect and quantify apoptosis of renal cells (37). Total cell number and apoptotic cells were counted first in the plane of the monolayer and then, by adjusting the focus, in the supernatant above. For each experimental condition, three randomly selected fields (×100 magnification) in each of three separate wells were counted. Etoposide (50 μM; Sigma, Poole, UK) was used as a positive control. Apoptosis was also demonstrated by Western blot for polyADP-ribose polymerase (PARP) cleavage. Nonadherent cells were collected by centrifugation, and pooled adherent and nonadherent cells were lysed using a gel-loading buffer comprising 62.5 mM Tris (pH 6.8), 6 M urea, 10% glycerol, 2% SDS, 0.0012% bromphenol blue, and 5% β-mercaptoethanol. Lysates were freeze-thawed (−80°C), sonicated, and denatured by boiling, and the proteins were resolved by SDS-polyacrylamide gel electrophoresis. Proteins were blotted onto nitrocellulose membranes, and PARP was detected with a monoclonal anti-PARP antiserum and secondary peroxidase-conjugated goat anti-mouse antibody. Bound antibodies were visualized using the ECL (Amersham, England) chemiluminescent visualization system.
DNA laddering characteristic of apoptosis was detected using ApoAlert LM-PCR assay kit (Clontech, Palo Alto, CA) according to the manufacturer’s instructions. Briefly, genomic DNA was extracted from pooled detached and adherent HPTC growing in 6-well plates under control or experimental conditions. A sample of 75 ng genomic DNA from each condition was used for adaptor ligation. An aliquot of 50 ng of adaptor-ligated DNA was used in a 100 μl volume of PCR reaction using 20 cycles. Then 15 μl of the reaction products were electrophoresed through 1.2% agarose gels containing 0.005% ethidium bromide at 75 V for 90 min. The DNA was visualized under ultraviolet light, and gels were photographed digitally.
Decoy Oligonucleotide Experiments
Decoy oligonucleotides were designed on the basis of the sequence of PPRE reported previously (38). The decoy (ACT TGA TCC CGT TTC AAC TC) or scrambled (TTA GGG AAT CAG CAA GAG GT) oligonucleotides were annealed to their respective complementary sequence by dissolving oligonucleotide pairs, each at a concentration of 100 μM, in a buffer containing 10 mM Tris-HCl (pH 7.5), 100 mM NaCl, and 1 mM EDTA. The oligonucleotide solution was warmed to 65°C in a water bath, maintained at 65°C for 10 min, and then cooled slowly (1 to 2 h) to room temperature. Stocks of annealed oligonucleotides were stored at −20°C. HPTC were transfected with 1 μM decoy or scrambled oligonucleotides together with pPPRE-TK-luc using Fugene 6 exactly as for other DNA constructs.
Assessment of Cell Number and Proliferation
Proliferation of cells after was determined after 24 h incubation with different potential PPAR agonists by [3H]-thymidine incorporation as described previously (7,8). In other experiments, cells growing in 12-well plates were treated with PPAR agonists for 24 h, washed with PBS, and trypsinized. Cell number for each condition was determined using a hemocytometer.
Statistical Analyses
Where appropriate, experimental data were analyzed by one-way ANOVA followed by either Dunnett or Tukey post hoc tests for multiple comparisons. P < 0.05 was considered significant.
Results
In preliminary experiments, expression of PPAR subtypes at the protein level in HPTC was examined by immunoblotting using specific antisera. PPARα, PPARβ/δ, and PPARγ were all found to be present in these cells (data not shown). Initial experiments with a variety of transfection reagents were also performed to determine the optimum conditions for the transient transfection of primary HPTC. Fugene 6 was found to be greatly superior to all other reagents tested. Judged by fluorescence microscopy of pEGFP-C1–transfected HPTC, a transfection efficiency of 50 to 60% was routinely achieved using the conditions described in Materials and Methods (results not shown).
In HPTC transiently transfected with pPPRE-TK-luc, 1 μM PGJ2, a PPARγ agonist, was able to stimulate a highly significant 5.4 ± 0.7–fold increase in luciferase expression compared with similarly transfected nonstimulated controls (Figure 1). The PPARα agonist ETYA evoked a lesser 1.9 ± 0.3–fold increase in luciferase expression. Interestingly, treatment with FA(+)HSA resulted in a dose-dependent increase in luciferase expression, maximal 2.4 ± 0.2-fold, at a concentration of 30 mg/ml (Figure 1). A similar effect was not observed with FA(−)HSA, indicating that bound FA were responsible for PPAR activation in HPTC.
Figure 1. Albumin-bound fatty acids are peroxisome proliferator–activated receptor (PPAR) response elements (PPRE) agonists in human proximal tubule cells (HPTC). HPTC were transiently transfected with pPPRE-TK-luc and pSV-βgal to normalize transfection efficiency. Transfected HPTC were treated for 24 h with the indicated concentrations of PGJ2, ETYA, FA(+), or FA(−) albumin. Cells were lysed, and luciferase reporter activity was assayed as described in Materials and Methods. Normalized luciferase activity is expressed as fold changes compared with control levels in nonstimulated cells. Results are expressed as means ± SEM of ≥5 experiments (* P < 0.05; ** P < 0.01 versus control).
HPTC were transfected with pCMX-RXRα and pSG5-PPARα or pCMX-PPARγ to achieve a twofold to threefold overexpression of the relevant PPAR (Figure 2). Transfection with pSG5-PPARδ resulted in approximately 1.5-fold overexpression of PPARδ (Figure 2). In cells overexpressing PPARγ, the ability of 1 μM PGJ2 to stimulate luciferase expression was augmented, being 11.4 ± 1.8–fold greater than controls (Figure 3). The luciferase response to ETYA, 2.2 ± 0.2–fold increase, was similar to that observed non-PPARγ–overexpressing cells. The luciferase response to FA(+)HSA was also significantly enhanced in PPARγ overexpressing cells, being maximal at a FA(+)HSA concentration of 50 mg/ml when luciferase expression increased 5.5 ± 1.2–fold compared with nonstimulated controls (Figure 3).
Figure 2. Expression of PPAR subtypes in nontransfected and transfected cells. Wild-type and PPAR-transfected HPTC were lysed and immunoblotted with specific anti-PPAR antisera. Expression levels of (A) PPARα, (B) PPARγ, and (C) PPARδ in nontransfected cells, lane 1, and PPAR-transfected cells, lane 2, are indicated by arrows. Blots are representative of several experiments.
Figure 3. Albumin-bound fatty acid stimulation of PPRE is augmented in PPARγ- but not PPARα-overexpressing cells. HPTC were transiently transfected with pPPRE-TK-luc, pCMX-PPARγ, or pSG5-PPARα, pCMX-RXRα, pSV-βgal to normalize transfection efficiency. Transiently transfected HPTC were treated for 24 h with the indicated concentrations of PGJ2, ETYA, FA(+), or FA(−) albumin. Cells were lysed, and luciferase activity in lysates was determined as described in Materials and Methods. Normalized luciferase activity is expressed as fold changes compared with control levels in nonstimulated cells. Results are expressed as means ± SEM of five experiments (* P < 0.05; ** P < 0.01 versus control).
In PPARα-overexpressing cells, the luciferase response to ETYA was enhanced (3.8 ± 0.8–fold) compared with non–PPARα-overexpressing cells. Conversely, overexpression of PPARα together with RXRα failed to augment luciferase expression in HPTC treated with PGJ2 or FA(+) HSA (Figure 3). Similarly, overexpression of PPARδ had no effect on PGJ2-, ETYA-, or FA(+)HAS-induced luciferase activity in HPTC (data not shown).
In preliminary experiments, we had observed that overnight incubation of HPTC with 10 μM or higher concentrations of PGJ2 led to extensive cell death. To determine the ability of PPARγ agonists to stimulate apoptosis, we studied HPTC morphology on exposure to these agonists. Nuclear morphology was examined using acridine orange, a technique previously used to detect apoptosis in renal cells (37). Representative images are depicted in Figure 4. Cells exposed to quiescence medium overnight display normal nuclear morphology (Figure 4A), but exposure of HPTC to similar medium containing the apoptosis-inducing agent, 50 μM etoposide, resulted in the appearance of typically shrunken apoptotic cells with contracted brightly staining nuclei (Figure 4B). When cells were exposed to either PGJ2 or FA(+)HSA at concentrations associated with strong activation of PPARγ, similar apoptotic cells appeared prominently (Figure 4, C and D). Conversely, when HPTC were exposed to FA(−)HAS, cells with apoptotic morphology were not easily detectable; indeed, mitoses were commonly observed (Figure 4E).
Figure 4. Induction of apoptotic nuclear morphology in nontransfected HPTC. HPTC growing in quiescence medium were treated for 24 h with the various indicated potential stimulators of apoptosis. Fluorescence microscopy of acridine orange–stained HPTC was used to study nuclear morphology of cells grown in quiescence medium alone (A), 50 μM etoposide (B), 1 mM PGJ2 (C), 30 mg/ml FA(+)HSA (D), or 30 mg/ml FA(−)HSA (E) by fluorescence microscopy after staining with acridine orange. Typical apoptotic cells are small with contracted brightly fluorescing nuclei. Photographs are representative of a least three individual experiments.
Examination of cell cultures by differential contrast interference microscopy (Normaski optics) revealed normal epithelial-appearing HPTC cultured in quiescence medium (Figure 5A). However, in the presence of PGJ2 and FA(+)HAS, shrunken rounded apoptotic-appearing cells were commonly seen (Figure 5, B and C). When incubated with 30 mg/ml FA(−)HAS, cells maintained a normal morphology (not shown).
Figure 5. Differential contrast interference microscopy reveals the presence of apoptotic HPTC after treatment with PGJ2 and FA(+)HSA. Nontransfected HPTC growing in quiescence medium were examined using Normaski optics under control conditions (A) or after 24 h treatment with 1 mM PGJ2 (B) or 10 mg/ml FA(+)HSA (C). Photographs are representative of a least three individual experiments. Apoptotic cells indicated by white arrows.
Further confirmation of induction of apoptosis was obtained by examining fragmentation of PARP by Western blotting in HPTC cultured under various conditions. Only intact PARP is found in floating and adherent HPTC cultured in quiescence medium alone or identical medium containing 30 mg/ml FA(−)HSA (Figure 6). In contrast, a typical apoptotic cleavage fragment of PARP was observed in HPTC cultured with PGJ2 and FA(+)HSA. A fainter band is present in the lane representing etoposide-treated cells. Floating and adherent cells cultured under various conditions were collected for extraction of DNA. The extracted DNA was examined for the presence of DNA laddering as described above. Typical apoptotic DNA laddering was detected in cultures of HPTC exposed to 50 μM etoposide, 1 μM PGJ2, and 30 mg/ml FA(+)HSA (Figure 7: lanes 3, 4, and 5, respectively). When HPTC were cultured in complete growth medium, quiescence medium, or in the presence of 30 mg/ml FA(−)HSA, apoptotic DNA laddering was not evident (Figure 6: lanes 1, 2, and 6, respectively).
Figure 6. Induction of poly(A)DP-ribose polymerase (PARP) cleavage by PPAR agonists in HPTC. Cells growing in quiescence medium were lysed under control conditions or after treatment with 1 μM PGJ2, 30 mg/ml FA(+)HSA, 30 mg/ml FA(−)HAS, or 50 μM etoposide for 24 h. Cleavage of PARP was examined by immunoblotting with specific anti-PARP antisera. The appearance of a PARP cleavage fragment is typical of apoptotic cells. The immunoblot depicted is representative of three identical experiments.
Figure 7. Induction of DNA laddering by albumin-bound fatty acids and PPARγ agonists. Laddering of DNA was detected as described in Materials and Methods in HPTC growing under control conditions in complete growth medium (lane 1) and quiescence medium (lane 2), or after 24 h of treatment with 50 μM etoposide (lane 3), 1 μM PGJ2 (lane 4), 30 mg/ml FA(+)HSA (lane 5), or 30 mg/ml FA(−)HSA (lane 6). The gel photograph depicted is representative of five experiments.
Apoptosis in HPTC was quantified by counting of typical nuclei stained by acridine orange according to the method of Mooney et al. (37). In nontransfected cells (Figure 8) under control conditions and cultured in quiescence medium, apoptotic cells were infrequent, comprising 2.2 ± 0.38% of the total. When treated with 50 μM etoposide, the number of apoptotic cells increased significantly to 8.3 ± 0.39%. Similarly, when cultured in the presence of either 1 μM PGJ2 or 30 mg/ml FA(+)HSA, apoptosis is increased significantly compared with controls at 15.5 ± 1.2% and 27.0 ± 1.1%, respectively. Conversely, treatment with 30 mg/ml FA(−)HSA failed to significantly increase apoptotic cell numbers compared with controls.
Figure 8. Albumin-bound fatty acids stimulate apoptosis of HPTC. Cells were grown in quiescence medium under control conditions or in the presence of the indicated concentrations of etoposide, PGJ2, FA(+)HSA, and FA(−)HSA. Acridine orange was added to culture medium, and apoptotic nuclei were counted with the cells in situ. Results are expressed as means ± SEM of three experiments (* P < 0.01).
Transient overexpression of RXR and PPARγ in HPTC results in an increase in apoptotic cells (12.1 ± 2.3%) compared with mock-transfected cells (2.0 ± 0.6%), even in cells cultured under control conditions (Figure 9). In mock-transfected HPTC incubated with 1 μM PGJ2, 30 mg/ml FA(+)HSA, or 30 mg/mlFA(−)HSA, levels of apoptosis were similar to those observed in nontransfected cells. However, in cells transfected with PPARγ and RXR, levels of apoptosis were markedly increased over those observed in mock-transfected cells for each experimental condition (Figure 9).
Figure 9. Overexpression of PPARγ enhances HPTC apoptosis induced by PGJ2 and albumin-bound fatty acids. HPTC were transfected with pCMX-PPARγ and pCMX-RXRα or mock transfected with empty vector. After transfection, cells cultured in quiescence medium were studied under control conditions or after treatment with PGJ2, FA(+)HSA, and FA(−)HSA at the indicated concentrations. Acridine orange was added to culture medium, and apoptotic nuclei were counted with the cells in situ. Results are expressed as means ± SEM of six experiments (* P < 0.05 versus mock control and P < 0.001 versus PPARγ transfected PGJ2–treated; ** P < 0.001 versus mock control and PPARγ transfected PGJ2–treated; *** P < 0.001 versus mock control, P < 0.001 versus PPARγ transfected PGJ2–treated, and P < 0.001 versus mock FA(−)HSA-treated; **** P < 0.01 versus mock control and P < 0.05 versus mock FA(−)HSA treated).
To inhibit PPAR function, cells were transfected with PPRE decoy or scrambled oligonucleotides (Figure 10). The transfection of these oligonucleotides had no effect on HPTC apoptosis under control conditions. Transfection of HPTC with scrambled oligonucleotides had no effect on levels of apoptosis induced by either 1 μM PGJ2 or 30 mg/ml FA(+)HSA, which were equivalent to levels seen in nontransfected cells at 42.0 ± 2.8% and 30.4 ± 8.72%, respectively. Decoy oligonucleotides, however, were able to significantly inhibit the apoptosis induced by both of these agents (Figure 10).
Figure 10. PPRE decoy oligonucleotides inhibit PGJ2 and albumin-bound fatty acid–induced apoptosis. HPTC were transfected with PPRE decoy oligonucleotides or scrambled (scram) control oligonucleotides. Transfected cells growing in quiescence medium were studied under control conditions or after treatment with PGJ2 or FA(+)HSA at the indicated concentrations for 24 h. Acridine orange was added to culture medium, and apoptotic nuclei were counted with the cells in situ. Results are expressed as means ± SEM of three or four experiments (** P < 0.01 versus decoy oligonucleotide transfcected PGJ2–treated; * P < 0.05 versus decoy oligonucleotide transfected FA(+)HSA–treated).
To confirm that decoy oligonucleotides were able to inhibit PPAR function, HPTC were transfected with pPPRE-TK-luc in the presence of scrambled or decoy oligonucleotides. As demonstrated in Figure 11, in the presence of scrambled oligonucleotides, 1 μM PGJ2 stimulated a 3.3 ± 0.9–fold increase in luciferase expression and 30 mg/ml (FA+)HSA a 2.2 ± 0.3–fold increase in luciferase expression, compared with similarly transfected controls cultured in quiescence medium. In the presence of decoy oligonucleotides, luciferase activity stimulated by both 1 μM PGJ2 and 30 mg/ml FA(+)HSA was completely attenuated (Figure 11).
Figure 11. PPRE decoy oligonucleotides inhibit activation of PPRE-driven luciferase expression by PGJ2 and FA(+)HSA. HPTC were transiently transfected with pPPRE-TK-luc, pSV-bgal to normalize transfection efficiency, and either PPRE decoy oligonucleotides or scrambled control oligonucleotides. Transfected HPTC were treated for 24 h with the indicated concentrations of PGJ2, ETYA, FA(+), or FA(−) albumin. Cells were lysed, and luciferase reporter activity was assayed as described in Materials and Methods. Normalized luciferase activity is expressed as fold changes compared with control levels in nonstimulated cells. Results are expressed as means ± SEM of three experiments (* P < 0.05 versus scrambled oligonucleotide-transfected control).
Proliferative responses of HPTC after treatment with PGJ2, FA(+)HSA, and FA(−)HSA were assessed in parallel with the apoptotic studies. Stimulation of PPARγ by PGJ2 resulted in a reduction in HPTC proliferation (0.39 ± 0.035-fold compared with control) as well as apoptosis, whereas, in contrast, FA(+)HSA was associated with both increased proliferation (4.2 ± 0.37-fold compared with control) and apoptosis (Figure 12A). Incubation of HPTC with FA(−)HSA resulted in a modest increase in proliferation as described previously (7,8). Cell numbers were markedly and significantly decreased in HPTC treated with PGJ2, and FA(+)HSA, but increased in the FA(−)HSA group (Figure 12B).
Figure 12. Proliferative responses of HPTC to agonist activation of PPARγ. (A) [3H]-thymidine incorporation into HPTC cultures in response to 24 h incubation with 1 μM PGJ2, FA(+), and FA(−)HSA. (B) Cell number determined by hemocytometer counting in HPTC cultures after 24 h incubation with 1 μM PGJ2, FA(+), and FA(−)HSA. Values are means ± SEM of three experiments (* P < 0.05 versus control; ** P < 0.01 versus control).
Discussion
Proteinuric renal disease is frequently associated with progressive tubular atrophy with interstitial inflammation and scarring. The end-stage kidney is remarkable for almost complete loss of proximal tubular elements and extensive tubulointerstitial fibrosis (39). Clearly therefore, the proteinuric environment may precipitate abnormalities of PTC growth and survival culminating in the development of the classical end-stage kidney. Studies of both human renal biopsy material and in vivo studies of animal models of renal disease have demonstrated alterations in PTC growth in proteinuria (4,5). These alterations are manifest as the simultaneous occurrence of both proliferation and apoptosis, with the balance appearing to favor apoptosis and thus eventual net deletion of cells.
The most abundant protein in proteinuric tubular fluid is albumin. Our previous work has demonstrated that in vitro albumin is able to stimulate proliferation of cultured PTC, and it seems likely that PTC proliferation observed in proteinuria in vivo may be elicited by albumin in tubular fluid (7,8). The precipitant of PTC apoptosis remains controversial.
Several investigators have examined the ability of large quantities of albumin to cause apoptosis in cultured PTC, but data on this issue conflict. Some authors have failed to demonstrate evidence of albumin-induced cytotoxicity in primary PTC cultures; indeed, other workers describe a survival factor–like effect of albumin in serum-starved primary PTC (12). Using an immortalized PTC line, one group has demonstrated induction of apoptosis by delipidated albumin after prolonged incubation (11).
Our current work with PPAR resolves the tension between these observations and ascribes both physiologic and pathophysiologic roles to PPAR in the human kidney. We postulated that in nephrosis albumin may act as a biologic Trojan horse, delivering injurious lipids to the interior of vulnerable PTC; we therefore chose to study the ability of albumin-bound lipids to stimulate lipid-activated transcription factors and thus modulate PTC growth and survival. Healthy albumin carries >99% of plasma FA (40), the FA content of plasma and albumin being very similar and comprising both saturated and unsaturated moieties (41). Data in nephrosis are scant (13, reviewed in reference 42); although the plasma albumin:FA ratio is greatly increased, the relative proportions of saturated and unsaturated species remain similar (42).
In nonminimal change disease, the urinary albumin FA load is indistinguishable from that of plasma albumin (13). Nephrotic proximal tubular fluid contains albumin concentrations approaching 3 mg/ml (43). The concentrations of albumin employed in the current experiments were thus higher than those likely to be encountered in the proximal tubule, even in heavy nephrosis. However our aim was to mimic the FA concentration prevailing in the proximal tubule in nephrosis, not that of albumin, using a pathophysiologically relevant human primary cell line. The FA(+)HSA used in these studies was derived from healthy humans; thus high albumin concentrations were selected to achieve total FA concentrations approximating those experienced in the nephrotic proximal tubule.
Our studies for the first time demonstrate that albumin-bound FA stimulate PPAR in a dose-dependent manner in HPTC. Augmented activation of a PPRE-luciferase reporter in PPARγ-overexpressing, but not PPARα or PPARδ–overexpressing, cells by PGJ2 and albumin-bound FA suggests that the PPAR activated under these conditions is indeed PPARγ. Treatment of HPTC with PPARγ agonists resulted in morphologic rounding of cells, condensation of nuclei, the cleavage of the caspase-3 substrate PARP, and laddering of DNA. These are the hallmarks of apoptosis; the results suggest that stimulation of PPARγ by PGJ2 and albumin-FA, but not albumin alone, results in apoptosis of HPTC. These results provide several lines of evidence supporting a critical role for PPARγ in the regulation of HPTC survival. First, PGJ2, a PPARγ agonist, potently induces PPAR transcriptional activation. Second, in the experiments where PPARγ was overexpressed, cell viability was reduced even in the absence of exogenous agonist. Third, double-stranded PPRE decoy oligonucleotides, but not a scrambled counterpart, inhibited both PPAR transcriptional responses to PGJ2 and albumin-FA and apoptosis induced by these two PPARγ stimulating agonists.
Opposing effects of PGJ2 and FA(+)HSA on cell proliferation were apparent from the [3H]-thymidine incorporation experiments. Whereas PGJ2 inhibited HPTC proliferation in association with apoptosis, pleiotropic effects of FA(+)HSA on cell growth and survival were observed, with the simultaneous occurrence of both apoptosis and proliferation. However, consideration of the relative fold changes in apoptosis and proliferation, together with cell counting suggested that the net effect of both of these agents is cell loss. Indeed, it is likely that cell counting overestimates the number of viable cells under these conditions with the inevitable inclusion of some apoptotic cells in the trypsinized cell suspensions used for hemocytometric analysis. The balance of apoptosis/proliferation observed in this in vitro study is reminiscent of that seen in our previous studies of protein overload nephrosis induced by either FA(+) albumin and FA(−) albumin (44). Overall, the results strongly support the notion of FA-evoked, PPARγ-mediated HPTC apoptosis, with an additional effect of albumin per se to stimulate cell proliferation.
Although data describing PPAR function in the kidney is virtually absent, some information relating to intrarenal localization of the three subtypes is available. Using RT-PCR of mRNA isolated from microdissected rat nephron segments, Yang et al. (26) found ubiquitous expression of PPARβ/δ in all segments examined. Expression of PPARα was restricted predominantly to the proximal convoluted tubule, whereas PPARγ was expressed in proximal straight tubule and inner medullary collecting duct. Using nuclease protection assays Guan et al. (25) demonstrated similar levels of expression of PPARβ/δ and PPARγ mRNAs in rabbit kidney cortex and medulla. Expression of PPARα mRNA predominated in the cortex. In contrast, only PPARα mRNA was detected in the cortex of rabbit and human kidney by in situ hybridization. Our studies are the first to examine expression of PPAR at the protein level in HPTC and indicate that all three PPAR subtypes are expressed in these cells. Unregulated exposure of HPTC to large quantities of albumin-bound lipid by an atypical apical route thus appears to precipitate activation of apoptosis-related pathways via PPARγ, although our results do not exclude the possibility that ligand activation of other PPAR may also induce apoptosis.
Results presented by several authors suggest a potential role for PPAR agonists in the treatment of renal disease (34). In animal models of diabetes, the PPARγ agonistic thiazolidinediones, as well as ameliorating metabolic derangements, protect against proteinuria, mesangial expansion, and glomerulosclerosis (45). These effects may be independent of the improvements in metabolic control, but they are rather related to direct effects of thiazolidinediones on glomerular cells. Asano et al. (46) reported inhibition of rat mesangial cell proliferation by troglitazone and PGJ2. Furthermore, Nicholas et al. (47) demonstrated that activation of PPARγ in cultured mesangial cells is associated with inhibited cell proliferation in a dose-dependent manner and suggested that activation of PPARγ may attenuate diabetic glomerular disease by blocking mesangial cell growth.
These drugs also appear to prevent glomerulosclerosis in the nondiabetic, rat remnant kidney model of renal disease. The salutary glomerular effects of thiazolidinediones in this model were associated with reduced glomerular cell proliferation, reduced expression of pro-fibrotic transforming growth factor-β, and lowered expression of plasminogen activator inhibitor-1 in the absence of improved BP control (48).
Do the profound effects of PPARγ activation on human PTC survival described in the current study represent a desirable or deleterious event? The results of our work certainly suggest that albumin-bound FA are tubulotoxic as a consequence of PPARγ activation. The findings of the current study are also supported by our recent observations that protein overload proteinuria induced in rats using FA(+) bovine serum albumin results in significantly greater tubular apoptosis and interstitial macrophage infiltration than is seen in animals rendered proteinuric by the injection of FA(−) bovine serum albumin (44).
However, clinical experience with thiazolidinediones in the treatment of type II diabetes does not suggest an increase risk of tubulointerstitial disease and progressive renal disease, at least as yet. Indeed animal studies suggest a protective effect of thiazolidinediones in both diabetic and nondiabetic models of renal disease (45–48). Rats and humans however exhibit different lipid profiles that make it difficult to directly extrapolate results form these animal studies to human disease. More studies in this area are required, along with greater focus on tubular pathology as opposed to glomerular consequences of renal disease.
Transcriptional activation of different gene sets by different PPAR agonists is ligand- and depot-specific (49–53). Even different members of the thiazolidinedione class of antidiabetic drugs acting via PPARγ induce differential gene transcription (50–52). Intense activation of PPARγ by PGJ2 and albumin-bound FA in PTC leads to apoptosis. Less-intense or thiazolidinedione-mediated PPARγ activation in the same cells may result in transcriptional activation of different, non–apoptosis-related gene sets. Alternatively, activation of proximal tubular PPARγ by thiazolidinediones in renal disease may inhibit potentially maladaptive proliferation of PTC short of induction of apoptosis. These issues require further investigation.
In summary therefore, albumin-bound FA are able to activate PPAR in HPTC. The consequences of this activation include apoptotic cell death. These studies confirm the injurious potential of filtered FA in nephrosis and provide further impetus for the use of lipid-lowering therapies in proteinuric renal disease.
Acknowledgments
Financial support for this work was provided by The Wellcome Trust and by project grant support from the National Kidney Research Fund.
- © 2003 American Society of Nephrology
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