* Division of Cell and Molecular Biology, Department of Biology, Boston University, 5 Cummington Street, Boston, Massachusetts 02215; 3M Medical Department, Corporate Toxicology, 3M Center 220-2E-02, Maplewood, Minnesota 55144; and
3M Corporate Research Analytical Laboratory, 3M Center 201-1W-29, Maplewood, Minnesota 55144
Received December 18, 2003; accepted March 8, 2004
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ABSTRACT |
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Key Words: perfluorooctanesulfonamide; perfluorooctanesulfonate; N-ethyl perfluorooctanesulfonamidoethanol; rodenthepatocarcinogenesis; ppar; peroxisome proliferation.
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INTRODUCTION |
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Peroxisome proliferation and certain other metabolic effects of PFOSAs may be a consequence of the activation of the nuclear receptor PPAR. PPAR
is a member of the nuclear receptor superfamily and mediates a broad range of biological responses to fatty acids, their synthetic analogs and structurally diverse lipophilic chemicals containing an acidic group (Gonzalez et al., 1998
). Peroxisomes are organelles involved in fatty acid oxidation, cholesterol metabolism, and hydrogen peroxide-linked respiration (Lock et al., 1989
). Activation of PPAR
leads to altered regulation of distinct sets of genes involved in lipid metabolism and homeostasis, peroxisome proliferation, and cell growth (Corton et al., 2000
). Long-term exposure of rodents to peroxisome proliferator chemicals causes a PPAR
-dependent increase in liver tumors (Peters et al., 1997
), raising concerns about the safety of these compounds for humans and other species (Cattley et al., 1998
). In contrast to rats and mice, guinea pigs and other species, including cynomolgus monkeys (Seacat et al., 2002
), show little or no evidence of peroxisome proliferation when treated with fluorochemicals or other peroxisome proliferating compounds. Studies with human hepatocytes suggest that humans are also likely to be poorly responsive to hepatic peroxisome proliferators. This species-dependence reflects several factors, one of which is the expression of PPAR
in humans and other unresponsive species at a substantially lower level than in rat and mouse liver (Palmer et al., 1998
).
A second important factor contributing to the species-specificity of peroxisome proliferative responses is the lower intrinsic responsiveness of human PPAR compared to rodent PPAR
to some, but not all, peroxisome proliferator chemicals (Keller et al., 1997
; Maloney and Waxman, 1999
). Human PPAR
requires a 5-fold higher concentration of the classical peroxisome proliferator Wy-14,643 for transcriptional activation when compared to mouse PPAR
activation (Keller et al., 1997
; Maloney and Waxman, 1999
). A similar species difference in PPAR
responsiveness is seen with the phthalate mono-ester mono(2-ethylhexyl)-phthalate (MEHP), a plasticizer metabolite and widespread environmental pollutant (Hurst and Waxman, 2003
). Human PPAR
also requires a higher concentration of perfluorooctanoate to achieve maximal activation than does mouse PPAR
(Maloney and Waxman, 1999
). Important differences in PPAR ligand specificity are also apparent in the comparison of PPAR
and PPAR
(Forman et al., 1997
). For example, PPAR
but not PPAR
is strongly activated by classic rodent peroxisome proliferators, such as Wy-14,643 and fibrate drugs (e.g., clofibrate, nafenopin), whereas PPAR
is preferentially activated by a distinct set of chemicals, including anti-diabetes type II drugs of the thiazolidinedione class, such as troglitazone. PPAR
can, however, be efficiently activated by certain activators of PPAR
, as shown in the case of MEHP (Hurst and Waxman, 2003
; Maloney and Waxman, 1999
). It is suspected that PPAR
can be activated by PFOSAs; however, it is uncertain whether species-dependent differences in receptor responsiveness, which could be important for human risk assessment, characterize this class of fluorochemicals.
The specific goals of the present study were (1) to ascertain whether, and with what potency, PFOSAs activate PPAR, the major mediator of hepatic peroxisome proliferation responses, as determined using trans-activation reporter assays and by evaluation of the responsiveness of endogenous PPAR
target genes in a liver cell model; and (2) to characterize the sensitivity of human PPAR
to PFOSAs, which may aid in the extrapolation to human toxicology and human risk assessment findings based on rodent model studies.
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MATERIALS AND METHODS |
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Fluorochemical analysis. Working solutions of each fluorochemical were prepared in DMEM as described below at concentrations of 8, 16, 32, 64, 125, 250, 500, and 1000 µM, with a final DMSO concentration of 0.1%. HPLC/electrospray mass spectrometry (ESI-MS) was used to quantify the concentrations of PFOS and FOSA in spiked DMEM samples using a Thermo-Finnigan LCQ HPLC/MS system (LCQ) and a Thermo-Finnigan aQa HPLC/MS system (aQa). Both HPLC/MS instruments included electrospray ionization sources that were operated in the negative ionization mode. The ESI probes were maintained at 4 kV and 285°C. LCQ mass spectral data were acquired by full scan with the selected-ion-current area of m/z 499 and m/z 498 being used for quantitation. aQa mass spectral data were acquired in the selected-ion-monitoring mode using the same masses for quantification. PFOS and FOSA were separated from other sample components using a 4 mm x 125 mm Phenosphere ODS (2), 5 µm column and a 30 to 95% acetonitrile gradient in water (over 4 min) at a flow rate of 0.5 ml/min. Both solvents contained 6 mM ammonium acetate as a modifier.
The responses of the HPLC/MS systems were calibrated by analyzing a series of PFOS and FOSA reference standards. The areas of the m/z 499 ions (PFOS) and the m/z 498 ions (FOSA) were used to generate calibration curves that were used to quantify the concentrations of PFOS and FOSA in the spiked MEM samples. The m/z 499 ion is the anion of PFOS acid and the m/z 498 ion is the (M-H) ion of FOSA. Prior to the analyses, each of the samples was diluted with acetonitrile such that the resulting solutions had PFOS and FOSA concentrations that produced responses within the linear response range of the HPLC/MS. Portions of these diluted solutions were then transferred to glass autosampler vials for HPLC/ESI-MS analysis. The areas of the m/z 499 and m/z 498 responses were compared to the calibration curves to calculate the PFOS and FOSA concentrations. The reported concentrations included the initial dilution factor in the concentration calculations.
N-EtFOSE concentrations in the spiked MEM samples were quantified using a Hewlett Packard Model 5890 Series II GC equipped with a flame ionization detector, a Hewlett Packard Model 7673 autosampler, and ChemStation data system. N-EtFOSE was separated from other compounds present in the samples using a Zebron ZB-5 30 meter fused silica capillary column having an inside diameter of 0.32 mm and a ZB-5 film thickness of 0.25 µm. Helium was used as the carrier gas at a flow rate of 40 cm/s. The column temperature was programmed from an initial temperature of 80°C (1-min hold time) to 190°C at a linear rate of 15°C/min, then to a final temperature of 340°C at a linear rate of 35°C/min (3-min hold time). A 2-µl portion of each diluted sample was injected using a 10:1 split ratio. Prior to sample analyses, the GC was calibrated with a series of N-EtFOSE reference standards.
Plasmids. The firefly luciferase reporter pHD(x3)Luc, obtained from Dr. J. Capone (McMaster University, Toronto, ON, Canada), contains three tandem copies of a PPAR-activated DNA response element derived from the rat enoyl-CoA hydratase/3-hydroxyacyl-CoA dehydrogenase gene linked directly to a minimal promoter (Zhang et al., 1993). Expression plasmids for mouse PPAR
and human PPAR
, respectively provided by Dr. E. Johnson (Scripps Research Institute, La Jolla, CA) and Dr. F. Gonzalez (National Cancer Institute, Bethesda, MD), were described previously (Hurst and Waxman, 2003
). Renilla luciferase reporter plasmid, pRL-CMV, was purchased from Promega (Madison WI). pBabe-puro expression vector containing the full-length cDNA encoding mouse PPAR
, which was used to generate FAO-PPAR
cells (see below), was obtained from Dr. B. Spiegelman (Dana-Farber Cancer Institute, Boston, MA).
Transient transfection assays. COS-1 cells were grown in Dulbecco's modified Eagles medium (DMEM) containing 10% fetal bovine serum. COS-1 cells were plated at a density of 3 x 104 cells/well of a 48-well tissue culture plate containing 500 µl of culture medium per well. The medium was replaced with 0.25 ml of fresh DMEM + FBS 24 h after plating, at which time transfection was carried out using 0.3 µl FuGENE 6 transfection reagent (Roche Diagnostics Corp., Indianapolis, IN) per well. Twenty-four h after addition of the DNA-FuGENE mixture to the cells, the medium was changed to serum-free DMEM containing each test compound at concentrations specified in each experiment. Cells were lysed 24 h later and Firefly luciferase and Renilla luciferase activities were measured using a dual luciferase assay kit (Promega). Transfections were performed using the following amounts of plasmid DNA/well unless indicated otherwise: 90 ng of pHD(x3)Lucxsxs, 5 ng of mPPAR or hPPAR
expression plasmid and 1 ng of pRL-CMV. Salmon sperm DNA was added to the plasmid DNA mix to give a total of 200 ng.
Cell treatment and data analysis. Fluorochemicals were dissolved in DMSO to give a 250 mM stock solution. Serial dilutions were prepared in DMSO, followed by a final 1000-fold dilution into serum-free DMEM. Stock solutions of fluorochemicals in DMSO were prepared in triplicate, followed by dilution of each sample into DMEM in duplicate, to give six replicates (tested in six parallel wells) for each fluorochemical at each concentration. Data presented are mean values ± SD based on n = 3 independent dilutions, with each value being an average of duplicate determinations, except as noted. Luciferase activity values were normalized for transfection efficiency by dividing the measured Firefly luciferase activity values by the Renilla luciferase activity obtained for the same cell extract, i.e., (Firefly/Renilla) x 1000. Statistical analysis was performed using SAS Jump software, Version 3. Individual pairwise comparison of means were performed using Student's t-test, with p < 0.05 deemed significant.
Construction of FAO cells that overexpress mPPAR. Rat hepatoma FAO cells (Moore and Weiss, 1982
) were obtained from Dr. J. Vanden Heuvel (Penn State University) and were grown in DMEM containing 5% fetal bovine serum. Transfection of the packaging cell line Bosc 23 with rat PPAR
-encoding retroviral plasmid DNA, harvesting of the retroviral supernatant, and infection of FAO cells were carried out as described previously (Jounaidi et al., 1998
). Pools of puromycin-resistant cells were selected using 2 µg/ml puromycin for 2 weeks. Drug-resistant clones (FAO-PPAR
cells) were selected and analyzed for nafenopin responsiveness by Western blot analysis of the PPAR
target gene peroxisomal 3-ketoacyl-CoA thiolase (PTL).
Western blotting. Cell extracts (40 µg) were electrophoresed on 10% Laemmli SDS gels, electrotransferred onto nitrocellulose membranes, and then probed with anti-acyl-CoA oxidase (ACOX), anti-PTL antibody, or anti-catalase, generously provided by Drs. T. Hashimoto and J. K. Reddy (Northwestern University, Chicago IL), as described elsewhere (Zhou et al., 2002). Antibody binding was visualized on x-ray film by enhanced chemiluminescence using the ECL kit from Amersham Pharmacia Biotech. Scans of Western blots were obtained using a Canon IX-4015 scanner and Ofoto software. Protein band intensities were quantitated using ImageQuant, v1.2 software (Molecular Dynamics, Sunnyvale, CA).
Quantitation of mRNA levels by real-time PCR. Relative cellular levels of rat 18S rRNA and PTL, ACOX, peroxisomal bifunctional enzyme (PBE), and urate oxidase mRNAs were quantified by real-time PCR analysis using the ABI 7900 Prism Sequence Detection System (Applied Biosystems). Total RNA was extracted using TRIZOL reagent (Gibco BRL) from FAO-PPAR cells that were seeded in six-well plates at 7 x 105 cells/well and treated for 48 h with nafenopin (100 µM) or the indicated fluorochemical beginning 4 h after cell seeding. The RNA obtained was treated with DNase I to remove contaminating DNA. SYBR Green real time PCR assays were used to quantify the following rat mRNAs: PTL (forward primer 5'-GGC-ACA-AGG-GCA-TCC-AAT-C-3', reverse primer 5'-GTG-CGC-TGT-CTT-TGG-TTC-AA-3'); ACOX (forward primer 5'-CCT-CTG-TCG-ACC-TTG-TTC-GG-3', reverse primer 5'-ACG-ACC-ACG-TAG-TGC-CAA-TG-3'); PBE (forward primer 5'-GCC-TTG-GGC-TGT-CAC-TAT-CG-3', reverse primer 5'-CAA-GCC-GAC-ACG-AGC-CTT-T-3'); urate oxidase (forward primer 5'-ACT-GCA-AGT-GGC-GCT-ACC-A-3', reverse primer 5'-CCC-AGG-TAG-CCT-CGA-AAT-CC-3'); and 18S rRNA (forward primer 5'-CGC-CGC-TAG-AGG-TGA-AAT-TC-3', reverse primer 5'-CCA-GTC-GGC-ATC-GTT-TAT-GG-3'). For reverse transcription reactions, 0.4 µg RNA was transcribed into cDNA using random hexamer primers and MuLV reverse transcriptase (Applied Biosystems). Each real-time PCR reaction contained SYBR Green PCR Master Mix and 300 nM of each primer in a volume of 4 µl and was carried out in triplicate. The PCR program was 50°C for 2 min, 95°C for 10 min, followed by 40 cycles of 95°C for 15 s, 60°C for 1 min and 95°C for 15 s. No PCR amplification was observed in control reactions that omitted reverse transcriptase or the cDNA template. Relative levels of PTL, ACOX, PBE, and urate oxidase mRNA were calculated for each cDNA sample after subtracting the threshold cycle (CT) for 18S RNA (determined in triplicate for each cDNA) from the CT values (determined in triplicate) for PTL, ACOX, PBE, and urate oxidase to adjust for small differences in the amount of cDNA template present in each sample (
CT). The average
CT for untreated FAO-PPAR
cells was then subtracted from the corresponding
CT for fluorochemical-treated cells (
CT) and the values were back-transformed (2
CT) to calculate the relative abundance of each RNA in the treated cells compared to untreated controls.
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RESULTS |
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FAO-PPAR cells were treated with PFOS, FOSA, or N-EtFOSE (150 µM) for 48 h. Western blot analysis revealed a substantial, dose-dependent induction of PTL protein (upper band of doublet; Fig. 5A) in FAO-PPAR
cells treated with either PFOS (panel A, lanes 36) or FOSA (panel A, lanes 710) compared to the DMSO control (panel A, lane 1). The fact that PFOS and FOSA both induced PTL expression indicates that these are true PPAR
-dependent responses. The clear, positive response of PTL to PFOS and FOSA contrasts with the weak response of ACOX (
2-fold) in FAO-PPAR
cell extracts after PFOS treatment (Fig. 5A). ACOX was induced 45-fold in FAO-PPAR
cells treated with nafenopin, which serves as a positive control for PPAR
activation (panel A, lane 2 vs. lane 1). N-EtFOSE had no reproducible effect on PTL protein levels in FAO-PPAR
cells (Fig. 5B), in agreement with the reporter gene assay presented in Figure 4. Weak induction of catalase (1.4-fold) was apparent in FAO-PPAR
cells after nafenopin treatment (Fig. 5A). Since catalase was nearly unchanged after PFOS and FOSA treatment (panel A, lanes 310), the catalase band intensities serve as a loading control.
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DISCUSSION |
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The activation of PPAR by PFOS is consistent with the hepatic peroxisome proliferative activity seen in a variety of studies where PFOS was administered to rodents over a relatively short time period. In one study, PFOS administered to male rats at 200 ppm (0.02% PFOS) in the diet for 2 weeks, resulting in a cumulative dose of
270 mg/kg, increased hepatic fatty ACOX activity
5-fold (Ikeda et al., 1987
). In another study, a 7-fold increase in fatty acid oxidation was observed in mouse liver after five days of dosing with PFOS (500 ppm in the diet,
125 mg/kg cumulative dose; Sohlenius et al., 1992
). In a third study, a
2-fold increase in lauroyl CoA oxidase activity was seen in male rats given a single ip injection of PFOS at 100 mg/kg, while hepatic peroxisome proliferative responses were weak or undetectable at comparable dosing levels of N-EtFOSE (Berthiaume and Wallace, 2002
). In contrast to these effects of PFOS in rats, no peroxisome proliferation responses are seen in guinea pigs given cumulative doses of 160 mg/kg PFOS or 640 mg/kg N-EtFOSE over four days by oral gavage (Wallace et al., 2001
). These findings in short-term assays are consistent with the report that PFOS is almost as potent a rodent peroxisome proliferator as the established peroxisome proliferator perfluorooctanoic acid (Sohlenius et al., 1992
).
The induction of peroxisome proliferation by PFOS in rats appears to exhibit a threshold dose response in short-term assays that rapidly reach relatively high tissue concentrations of PFOS. A threshold dose-response would be consistent with the hypothesis that peroxisome proliferation is a receptor-dependent mechanism that exhibits nonlinearity and a threshold exposure requirement for gene induction responses (Lapinskas and Corton, 1997). In contrast, longer-term repeat-dose studies that achieve higher liver PFOS concentrations than necessary to induce peroxisome proliferation in a short-term assay show no significant increase in hepatic peroxisome proliferative response. For example, at four weeks into a chronic (two-year) PFOS bioassay, rats fed 20 ppm PFOS in the diet received cumulative doses of
52 and 50 mg PFOS/kg with corresponding serum PFOS concentrations of
42 ppm (
84 µM) and
54 ppm (
108 µM), in male and female rats, respectively (Seacat et al., 2003
). These serum concentrations were associated with
7-fold higher liver PFOS concentrations, 282 ppm and 373 ppm, respectively, indicating liver accumulation. However, only a moderate (
2-fold) increase in hepatic palmitoyl CoA oxidase activity was observed in the male liver samples and no response was seen in female liver samples, suggesting a very weak peroxisome proliferative response. Indeed, after 14 weeks of dosing, PFOS concentrations were greater than 130 ppm (
260 µM) in the serum and 450 ppm (
900 µM) in the liver with no increase in palmitoyl CoA oxidase activity (Seacat et al., 2003
). The modest increase in palmitoyl CoA oxidase activity seen in these studies suggests that the cumulative doses after 4 or 14 weeks of 20 ppm dietary PFOS may not have been achieved rapidly enough to produce the increases in palmitoyl CoA oxidase seen in the shorter-term exposure studies, despite similar cumulative doses. The presence of high tissue concentrations of PFOS without a sustained increase in palmitoyl CoA oxidase activity suggests that a mechanism may exist in vivo for an adaptive down-regulation of the hepatic peroxisome proliferation response to PFOS treatment, which could explain the low level of ACOX induction seen in the current in vitro study. Alternatively, the current ACOX results may not reflect what occurs in vivo and may occur by some other mechanism.
The present findings establish the responsiveness of mouse and human PPAR to perfluorooctane-based chemicals, and support other data indicating a role for PPAR
as a mediator of some of the toxicities associated with PPAR
activation, at least in rodent species. PFOS activation of PPAR
was characterized by an EC50 of 1315 µM in the luciferase reporter gene assay used in the present study. This concentration corresponds to a serum PFOS concentration of
58 ppm, or approximately 1220% of the in vivo serum PFOS concentration that elicited only a weak peroxisome response in male rat liver after 4 weeks administration of 20 ppm dietary PFOS (Seacat et al., 2003
). Other mechanisms for PFOS-based toxicity are likely, given the other cellular responses activated by these compounds in vitro, such as the inhibition of mitochondrial bioenergetics (Starkov and Wallace, 2002
) and effects on cell-cell communication via gap junctions, which occur in rat liver at PFOS concentrations ranging from 112 ppm to 810 ppm, without a concentration-dependent response in that range (Hu et al., 2002
). Presumably, these effects of PFOS involve mechanisms that are independent of PPAR
.
Some inconsistencies and variations in the dose-response data for PPAR activation were encountered in the present study, in particular for FOSA. This variability made it difficult to establish precise dose-response curves for receptor activation by FOSA. Nevertheless, the present studies establish the capacity of this fluorochemical for PPAR
activation, which is associated with hepatocarcinogenesis in rodents (Gonzalez et al., 1998
). Additional investigation established that the PPAR
-activation capacity of FOSA, and PFOS, extends to the induction of endogenous PPAR
target genes, as shown by the strong increase in expression of the peroxisomal enzymes PTL and PBE in FAO rat hepatoma cells that stably express PPAR
. Interestingly, PFOS and FOSA stimulated only a low level increase in expression of ACOX expression in FAO-PPAR
cells, even though this peroxisomal enzyme, a known PPAR
target gene, was readily induced by treatment of the cells with the established PPAR
ligand and trans-activator nafenopin. This indicates that ACOX activity, commonly used to monitor hepatic peroxisome proliferation, may be a relatively insensitive monitor of the effect of PFOS in rodent liver. Conceivably, PFOS and FOSA could bind to PPAR
in a manner that leads to trans-activation of a subset of classic PPAR
target genes, perhaps inducing a novel metabolic profile in terms of hepatic peroxisomal enzyme expression. Additional studies, including more detailed enzymatic analysis and global mRNA expression profiling of livers obtained from PFOS-treated rats and mice, may help clarify this issue. Some inconsistencies in the concentration of PFOS and FOSA required to activate a PPAR
luciferase reporter, induce mRNA and protein of PPAR
-responsive genes were seen. These may be explained by interexperiment variability due to fluorochemical solubility issues, which were routinely encountered in this study. Although studies in human cell lines would further characterize the risks posed by exposure to fluorooctane-based chemicals, we expect that such experiments would yield a negative result, given extensive earlier studies demonstrating the unresponsiveness of human cells to PPAR
activators (Cattley et al., 1998
; Lake, 1995
; Roglans et al., 2002
).
Peroxisome proliferators and other chemicals may activate PPARs by two types of mechanisms: by binding directly to the receptor (i.e., as ligands), and by perturbing lipid metabolism and transport in a manner that stimulates the synthesis and/or release of endogenous PPAR ligands. We cannot distinguish between these two mechanisms in the case of the activation of PPAR by PFOS and FOSA. Previous investigations have shown, however, that PFOSAs can displace endogenous ligands from liver fatty acid binding protein (Luebker et al., 2002
), an intracellular lipid carrier that binds and transports fatty acids, acyl-CoA derivatives, and a variety of other hydrophobic molecules within hepatocytes. This fatty acid binding protein stimulates phospholipid synthesis and regulates lipid metabolism, and may protect the cell by maintaining intracellular free fatty acids at subtoxic concentrations. Consequently, the hypolipidemic and peroxisome proliferating effects of PFOSAs may, in part, be due to interference with the capacity of liver fatty acid binding protein to bind fatty acids, cholesterol, and other lipids. Displacement of fatty acids could, in turn, stimulate the activation of PPAR
or PPAR
by these endogenous ligands. Further investigation, including a more direct examination of the ability of PFOSAs to bind directly PPAR
should help resolve this issue.
Finally, extension of the present findings to include other PPAR forms would be of interest, in view of their importance in diverse physiological processes including adipogenesis (PPAR; Schoonjans et al., 1996
), reverse cholesterol transport (Oliver et al., 2001
) and development (Peters et al., 2000
; PPAR
). In contrast to PPAR
, which is expressed at comparatively low levels in human cells (Palmer et al., 1998
), PPAR
is expressed at high levels in a broad range of human tissues. These include adipose tissue, where many lipophilic foreign chemicals tend to accumulate, as well as colon, heart, liver, testis, spleen, and hematopoietic cells (Vidal-Puig et al., 1997
). Other biological responses linked to PPAR
ligands and activators, which may potentially be influenced by fluorochemicals that activate PPAR
, include colon tumorigenesis (Saez et al., 1998
) and the inhibition of human vascular endothelial cell differentiation and angiogenesis (Xin et al., 1999
). PPAR
activation also leads to foam cell macrophage differentiation, although it is uncertain whether this latter PPAR
-dependent response is likely to contribute to generation of an atherosclerotic plaque (Rosen and Spiegelman, 2000
). The extent to which PFOSAs alter these physiological and pathophysiological responses to endogenous or other foreign chemical PPAR
ligands is uncertain, and is an important area for further research.
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ACKNOWLEDGMENTS |
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NOTES |
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1 To whom correspondence should be addressed. David J. Waxman Fax: (617) 353-7404. E-mail: djw{at}bu.edu.
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