Department of Pharmaceutics, Ernest Mario School of Pharmacy, Rutgers, The State University of New Jersey, 160 Frelinghuysen Road, Piscataway, New Jersey 08854
1 To whom correspondence should be addressed at Department of Pharmaceutics, Ernest Mario School of Pharmacy, Rutgers, The State University of New Jersey, 160 Frelinghuysen Road, Piscataway, NJ 088548022. Fax: (732) 445-3134. E-mail: gknipp{at}rci.rutgers.edu.
Received September 29, 2004; accepted December 15, 2004
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ABSTRACT |
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Key Words: phthalates; rat; placenta; HRP-1; fatty acid; transport; PPAR.
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INTRODUCTION |
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Human exposure to DEHP may begin in the mother's womb, where DEHP has been demonstrated to readily cross the placenta and accumulate in the fetus (Kihlstrom, 1983; Singh et al., 1975
). While the pathological consequences of DEHP exposure in human are uncertain, DEHP is an established reproductive and developmental toxicant and hepatocarcinogen in rodents, where respiratory distress and developmental disorders in fetal/neonatal testis, liver, and other major organs have been observed (Cammack et al., 2003
; Kavlock et al., 2002
; Magliozzi et al., 2003
; Moore et al., 2001
). DEHP, MEHP, and EHA belong to a diverse class of peroxisome proliferator chemicals (PPCs). Administration of PPCs to rodents results in a pleiotropic response characterized by peroxisome proliferation as well as numerous alterations in gene translation of proteins involved in essential fatty acid (EFA) transport, metabolism, and lipid homeostasis (Lemberger et al., 1996
; Simpson, 1997
). Such effects have been identified to be mediated by the peroxisome proliferator-activated receptors (PPARs), established regulators of cellular EFA homeostasis (Lemberger et al., 1996
). Three PPAR isoforms (
, ß, and
) have been identified in various tissues including placenta (Lemberger et al., 1996
; Wang et al., 2002
). They regulate the transcription of target genes by binding to PPAR response elements (PPRE) as heterodimers with retinoic X receptors (Bocher et al., 2002
; Lemberger et al., 1996
). PPAR
has been demonstrated to play a role in regulating lipid catabolism, whereas PPAR
controls adipocyte differentiation and lipid storage (Bocher et al., 2002
; Lemberger et al., 1996
). Although PPARß is less understood, it might be a mediator in the control of brain lipid metabolism, fatty acid induced adipogenesis, and atherogenic inflammation (Bocher et al., 2002
).
Fatty acids, especially EFAs and their long chain polyunsaturated fatty acids derivatives (LCPUFAs), are involved in energy storage and serve as obligatory constituents of biological membranes and precursors of intra- and intercellular signaling molecules in the body (Innis, 2003; Uauy et al., 1999
). Mammals are unable to synthesize EFAs, and therefore, the fetus relies on maternal dietary intake of EFAs and their directional placental transfer to obtain sufficient quantities for normal development (Dutta-Roy, 2000
; Haggarty, 2002
). An active, directional mechanism mediated by several distinct fatty acid transporters has been suggested to contribute to this directionality, which should compensate for the limited transfer capacity by simple free diffusion (Dutta-Roy, 2000
; Haggarty, 2002
). These fatty acid transfer-conferring proteins have been found in the rat and human placentas as well as in vitro trophoblast models (Dutta-Roy, 2000
; Haggarty, 2002
; Knipp et al., 1999
, 2000
), and include fatty acid transport protein 1 (FATP1), plasma membrane fatty acid binding protein (FABPpm), and heart cytoplasmic fatty acid binding protein (HFABP).
It is not clear how fetal exposure to DEHP and its metabolites leads to subsequent health hazards. Therefore, the present study was conducted to assess if the teratogenicity of these xenobiotics is related to placental handling of EFAs via PPAR trans-activation. The HRP-1 rat trophoblast model was utilized, which exhibits characteristics resembling those of the trophoblasts in the labyrinthine zone (rat placenta transport barrier) (Soares et al., 1987). HRP-1 cells have been previously demonstrated to form a monolayer of viable, polarized, fully differentiated cells and to be a useful in vitro model of the placental barrier to study transport of nutrients, including fatty acids (Shi et al., 1997
), glutamate (Novak et al., 2001
), and glucose (Das et al., 1998
). In this study, the effects of DEHP, MEHP, and EHA on the expression of the three PPAR isoforms (
, ß, and
), FATP1, FABPpm, and HFABP were investigated, as well as the functional influences of these compounds on the uptake and transport of long chain fatty acids.
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MATERIALS AND METHODS |
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Cell culture.
The HRP-1 trophoblastic cell line was a generous gift from Dr. Michael J. Soares (University of Kansas Medical Center, Kansas City, KS) and was cultured as described previously (Knipp et al., 2000). The 24-well culture plates and polycarbonate Transwell clusters were coated with type I rat tail collagen (Bio-Rad) at 5 µg/cm2 on the same day of cell seeding. For the expression studies, the HRP-1 trophoblastic cells were seeded at 5 x 104 cells/cm2 in T-25 cm2 flasks. For the uptake studies, the HRP-1 trophoblastic cells were seeded at 2.5 x 104 cells/cm2 in 24-well tissue culture plates. For the transport studies, the cells were seeded at 7.5 x 104 cells/cm2 on Transwell 12-well-clusters. The medium was aspirated and replaced with fresh medium every day. The expression/uptake or the transport studies were performed when the cells came to 90% to 95% confluence. All cells used in these studies were between passages 15 and 25.
Time- and dose-dependent exposure to xenobiotics.
Confluent HRP-1 cells were treated with DEHP, MEHP, and EHA diluted 1000-fold in assay buffer from a stock solution in DMSO at the indicated concentrations and harvested at the respective time periods for total RNA isolation and/or whole protein extraction as well as for the uptake and transport studies. Cells treated with 0.1% DMSO were used as negative control.
For the time-dependent study, HRP-1 cells were treated with 50 µM of DEHP, MEHP, and EHA and harvested at 2, 4, 8, 12, and 24 h. For the dose-dependent study, HRP-1 cells were treated with DEHP, MEHP, and EHA at the concentrations of 25, 50, 100, and 200 µM and harvested at 4 h and 12 h for mRNA and protein expression assays, respectively.
Semiquantitative RT-PCR.
Total RNA isolation, RT-PCR, and gel electrophoresis were performed as described previously (Wang et al., 2002) with optimized conditions and normalized to ß-actin expression using gene-specific primers (Table 1). PCR reaction products were electrophoretically separated with a 1.5% agarose gel. Ethidium bromide stained bands were visualized, and the resulting densitometry analysis was performed using a NucleoTech 920 Image detection system (NucleoTech Corporation, San Mateo, CA). The molecular weight for each band was determined in reference to a 100 bp ladder.
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Protein analysis.
Western and immunoblotting analyses were performed to check the specificity of the antibodies and to quantitate protein expression levels according to the manufacturer's protocol (Bio-Rad), respectively. Briefly, 30 µg of protein was loaded per lane or per well. In Western blotting experiments, the protein lysates were separated in 8 to 15% SDS polyacrymlamide gels under reducing conditions and transferred to nitrocellulose membrane (Millipore, Bedford, MA). For immunoblotting, the protein lysates were loaded into the well and allowed to sit for 10 min; the lysate was then transferred to the nitrocellulose membrane under vacuum. The resulting membranes were handled in the same manner by first blocking with 5% nonfat milk and then incubating overnight with the respective primary antibody (1:500 to 1:2000 with respect to each antibody, as described previously (Knipp et al., 2000; Wang et al., 2002
). The membranes were then washed 3 x 7 min with TBS/T (1x TBS containing 0.05% Tween 20) and then incubated for 1 h with a 1:5000 dilution of the appropriate HRP-conjugated secondary antibodies (Sigma Aldrich). The specific protein bands were visualized using chemiluminescence detection with the Pierce Femto Signal Western Reagent kit and recorded by the NucleoTech 920 Image detection system (NucleoTech Corporation, San Mateo, CA). To normalize the signal for these proteins, ß-actin levels were probed as internal controls.
Uptake assays.
Both the uptake and transport studies were performed in Hank's balanced salt solution (HBSS, Mediatech Inc.) containing 136.7 mM NaCl, 4.167 mM NaHCO3, 0.385 mM Na2HPO4, 0.441 mM KH2PO4, 0.952 mM CaCl2, 5.36 mM KCl, 0.812 mM MgSO4, 5.5 mM D-glucose) supplemented with 10 mM HEPES (pH 7.4). The stop buffer was prepared with 0.1% bovine serum albumin (BSA, Sigma Chemical Co.) and 200 µM phloretin (Sigma Chemical Co.) diluted in uptake/transport buffer (Xu et al., in press). The uptake and transport assay buffers were freshly prepared and comprised of HBSS with 200 µM of different LCFA (fatty acid: BSA molar ratio = 1:1) containing 0.1 µCi/ml of the appropriate 14C-radiolabeled LCFA, as previously described (Campbell et al., 1997; Xu et al., in press). The uptake study was performed for 15 min and stopped by rapidly washing the cells three times with ice-cold stop buffer (750 µl/well) followed by one rinse with ice-cold PBS (750 µl/well). The cells were then solubilized with 0.2 N NaOH (250 µl/well) and neutralized with 0.2 N HCL (250 µl/well). An aliquot of 425 µl was removed for scintillation counting, and a 25 µl aliquot for the BCA protein assay.
Transport Assays
Confirming monolayer integrity.
HRP-1 cell monolayer integrity was evaluated with trans-epithelial electrical resistance (TEER) measurements and mannitol permeability studies. The TEER values of the cell monolayers were determined by electrical resistance measurements at 37°C, using the Epithelial Voltohmmeter (EVOM, World Precision Instruments, Sarasota, FL) and corrected by the values obtained across the blank collagen-coated Transwell. Permeability studies of mannitol were performed in the influx (i.e., apical to basolateral) and efflux (i.e., basolateral to apical) directions to check the cell monolayer integrity.
Long chain fatty acid transport assays.
Transport studies were performed in the influx (AB, AB) and secretory efflux (B
A, BA) directions with the respective 14C-labeled fatty acid and 3H-mannitol, the latter as a marker for monolayer integrity as previously described (Xu et al., in press). For all of the studies, the apical and basolateral test solution volumes were 0.5 and 1.5 ml, respectively, and maintained at 37°C. Each cell monolayer used in the studies was rinsed and equilibrated for 30 min with the transport buffer. Following the addition of the donor solution, samples from the receiver side were removed at 15, 30, 45, 60, 75, 90, and 120 min and replaced with an equal volume of prewarmed transport buffer. The apical and basolateral sampling volumes were 50 µl and 150 µl, respectively, and all sample concentrations were corrected for dilution factors. At the end of the study, 50 µl of the donor side sample was collected to determine mass balance.
Data analysis.
All experiments were repeated a minimum of three times. Data generated from RT-PCR and slot blot were quantitated by densitometry (Gel ExpertTM software program, NucleoTech Corp., San Mateo, CA) and normalized to ß-actin expression.
The apparent permeability coefficients (Papp) were calculated using Equation 1:
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The permeability coefficients for the cell monolayer, Pmono, were calculated using Equation 2:
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All the data were presented as means ± standard deviation (SD) of at least three replicates. Statistical significance relative to vehicle controls, shown in each figure, was determined using a one-way ANOVA followed by the Student t-test, where p values <0.05 were considered to be statistically significant.
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RESULTS |
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Consistent with the mRNA studies, protein expression of PPAR and PPAR
increased with xenobiotic exposure when contrasted with the control cells, reaching statistical significance at 50 µM concentration of each reagent at 8 h (for PPAR
, p < 0.001) (Fig. 2, Panel C2) and 12 h (for PPAR
, p = 0.005) (Fig. 2, Panel A2). Similar to the dose-dependent induction patterns observed in the mRNA experiments, PPAR
and PPAR
protein expression were highest at 50 or 100 µM (Fig. 2 and 3, Panel A2 and C2).
The mRNA expression of PPARß was significantly induced at 2 and 4 h (p < 0.05) (Fig. 2, Panel B1) at the concentration of 50 µM for MEHP and DEHP, respectively. However, the induction did not result in a concurrent statistically significant increase in protein expression (Fig. 2, Panel B2) with one exception. At 12 h with 100 µM MEHP, PPARß protein expression had significantly increased versus controls (1.4-fold induction, p < 0.05) (Fig. 3, Panel B2). Thus, despite the increase observed at several time/dose points, PPARß seemed to be largely unchanged.
Effects of DEHP, MEHP, and EHA on the Expression of FATP1, FABPpm, and HFABP
As illustrated in Panel A1 of Figure 4, exposure to 50 µM of DEHP and its metabolites had dramatic effects on the mRNA expression of FATP1 at 4 h, and to a slightly less extent at 8 h (Fig. 4, Panel A1). The initial induction time points were generally later than those observed for PPAR and PPAR
(2 or 4 h). The peak FATP1 mRNA levels noted for DEHP (2.6-fold, p < 0.001) and MEHP (2.6-fold, p < 0.01) at 4 h were higher in contrast to EHA (1.75-fold, p < 0.05). FATP1 mRNA expression declined at 24 h upon exposure to 50 µM EHA (p < 0.05), which was similar to the effects observed with PPAR
and PPAR
(Fig. 2, Panel A1 and C1). FATP1 protein expression reached statistically significant induction levels at 12 and 24 h (p < 0.05), although levels appeared to trend upwards since 4 h. Obvious dose-dependent changes in response to the xenobiotics were determined, with peak levels at 50 and 100 µM for both mRNA (p < 0.05) (Fig. 5, panel A1) and protein (p < 0.05) (Fig. 5, panel A2) expressions of FATP1.
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Effects of DEHP, MEHP, and EHA on the Uptake of Long Chain Fatty Acid
To confirm that a concurrent functional change was occurring with the observed mRNA and protein changes, functional uptake/transport studies were performed after 24-h exposure to 50 µM DEHP, MEHP, and EHA, respectively. In addition, 100 µM of fenofibrate, a well-established PPAR agonist (Guay, 2002
) was used as a positive control, and 0.01% DMSO as negative control. Several representative LCFAs including AA (20:4,
-6), DHA (22:6,
-3), LA (18:2,
-6), ALA (18:3,
-3), OA (18:1,
-9) and SA (18:0, saturated) were selected for these studies. Time-dependent uptake studies revealed that each of the respective LCFAs demonstrated linear uptake kinetics until 30 min, which led to the selection of 15 min for the subsequent uptake studies (data not shown). The physiological, nonesterified fatty-acid-to-albumin ratio found in the human maternal plasma at term is about 1.3:1, and one BSA molecule has the capacity to binding up to six LCFA molecules (Benassayag et al., 1997
). The subsequent experiments were performed with a 1:1 molar ratio of fatty acid to BSA.
Consistent with previous reports (Campbell et al., 1997; Dutta-Roy, 2000
), the uptake rates of the polyunsaturated fatty acids (LA and ALA) were higher than those of the saturated (SA) and monounsaturated (OA) fatty acids of the same chain length (p < 0.05 for all comparisons) (Fig. 6). For the
-6 series of AA and LA, the uptakes rates decreased significantly (p < 0.01) by 65.5% when the chain length increased from C18 (LA) to C20 (AA). A similar trend was observed for the
-3 series, where the uptakes rates of DHA (C22) decreased 63% (p < 0.01) compared with ALA (C18) (Fig. 6). In addition, the uptake rate of AA was statistically higher than that of DHA (p < 0.01), with a similar phenomenon observed for LA versus ALA (p < 0.05) (Fig. 6). These results suggest that the
-6 family (AA, LA) demonstrated higher uptake rates than
-3 family (DHA, ALA) in these untreated samples.
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Effects of DEHP, MEHP, and EHA on the Transport of LCFA
TEER values and mannitol permeability were utilized to confirm cell monolayer integrity. In our preliminary studies, TEER increased to a maximum of 50 to 60 ·cm2 at 4 to 5 days post-seeding and then leveled off (data not shown). Thus, 50
·cm2 was chosen as the critical point of cell integrity and selection of the Transwells for subsequent fatty acid transport studies. Mannitol permeability was also investigated in both the influx and efflux directions. To further ensure the cell integrity upon exposure to phthalates, all of our fatty acid transport studies were performed with 14C-fatty acid and 3H-mannitol. The mannitol permeabilities (35 x 105 cm/s) did not change under the exposure to DEHP, MEHP, and EHA.
AA and DHA were selected as representative of the -3 and
-6 EFAs. Since they are polyunsaturated EFAs, the possibilities of appearance of metabolites in the donor and receiver chambers were investigated at the end of the transport study by reverse-phase high performance liquid chromatography (HPLC) in our preliminary studies. The fact that no metabolites were found (data not shown) excluded the potential disturbance of EFA metabolism from transport study. In addition, the mass balance matched very well (95 to 99%).
Although the influx (Pmono,influx) and efflux (Pmono,efflux) permeabilities of DHA were higher than those of AA in the untreated control cells, neither of them demonstrated obvious directional transport, with influx permeability ratios (Pmono,influx/Pmono,efflux) of 0.90 ± 0.07 and 1.03 ± 0.39 for AA and DHA, respectively. However, upon treatment with DEHP and its metabolites, the influx permeability ratios of AA and DHA were differentially induced (Fig. 7), although Pmono,influx and Pmono,efflux of AA and DHA were both elevated (data not shown). Under the exposure to the metabolites of MEHP and EHA, the influx ratio of AA was elevated significantly relative to the control cells, with 13.2 (p < 0.01) and 7.9% (p < 0.001) induction, respectively. DEHP treatment also seemed to slightly induce the influx ratio of AA (7.7%), but did not reach statistical significance (Fig. 7, Panel A). However, with DHA, DEHP elicited a significant increase in the influx ratio (76%, p < 0.01) (Fig. 7, Panel B).
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DISCUSSION |
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The concentrations of xenobiotics used in this study were in the range of 25 to 200 µM (9.76 to 78.1 µg/ml). This range is higher than what was observed in human maternal plasma at term in health subjects, where the mean DEHP and MEHP concentrations were 1.15 ± 0.81 and 2.05 ± 1.47 µg/ml, respectively (Latini et al., 2003). However, patients having a long-term exposure to DEHP-containing devices were reported to have a whole blood DEHP concentration of about 70 to 80 µg/ml, and neonatal exposure to DEHP following exchange transfusion with PVC catheters could increase blood levels to the range of 13.2 to 84.9 µg/ml (Kavlock et al., 2002
). Furthermore, to correlate toxic effects observed in animals with acceptable levels of exposure for humans, 10-fold increases in DEHP exposure are commonly used for the inter- and intraspecies variability.
Differences in DEHP toxicities have been demonstrated to be organ/tissue and species specific. It is well understood that DEHP produces a range of hepatic effects, which is believed due to the activation of PPAR (Kavlock et al., 2002
). However, the testicular, renal, developmental, and other extrahepatic toxicities exhibited by DEHP have been demonstrated to be independent of PPAR
, as suggested from the PPAR
knockout mouse studies (Peters et al., 1997
). Considering that the metabolites of DEHP, especially MEHP, activate both PPAR
and PPAR
in extrahepatic cell lines and/or tissues and is a stronger teratogen in mice (Kavlock et al., 2002
; Lampen et al., 2003
; Maloney et al., 1999
), it is feasible that DEHP acts through its metabolites to activate a PPAR-mediated signaling pathway. In accordance with this concept, our experiments did demonstrate an induction of PPAR
and PPAR
in a time- and dose-dependent manner (Figs. 2 and 3, Panel A and C) in HRP-1 cells. Furthermore, a relatively higher dose and longer exposure period of DEHP was needed compared to MEHP and EHA to obtain the same level of gene/protein activation. It should be noted that the responses of PPAR
and PPAR
to DEHP, MEHP, and EHA were similar, but not exactly concordant, which suggests different activities of the two isoforms in placental EFA transport/metabolism. PPAR
knockouts are known to be embryonic lethal between day 9.5 and 10.5 of gestation due to improper placentation (Barak et al., 1999
), suggesting the importance of this isoform in regulating fetal development.
There was no consistently significant pattern of PPARß expression regulation by DEHP, MEHP, and EHA (Figs. 2 and 3, Panel B), which is in accordance with previous reports (Kavlock et al., 2002). However, a recent investigation by Lampen et al. (2003)
demonstrated that MEHP and EHA activated mouse and human PPARß as well as PPAR
and PPAR
in an embryonic stem cell line. Actually, PPARß was also essential in placentation, because its deficiency resulted in frequent embryonic lethality (Barak et al., 2002
). Further study is needed to elucidate the regulation of PPARß by phthalates and other xenobiotics.
FATP1 and FABPpm are cell surface proteins involved in placental fatty acid uptake and transport (Knipp et al., 1999, 2000
), This study suggests that the mRNA and protein expression of FABPpm did not change upon administration of DEHP and its metabolites (Figs. 4 and 5, Panel B), which is consistent with a lack of PPAR regulation of FABPpm reported in the literature (Motojima et al., 1998
). Considering that FABPpm is abundantly expressed throughout the rat chorioallantoic placenta, it probably plays a generalized compensatory function in facilitating fetal fatty acid uptake (Knipp et al., 1999
, 2000
). Although Motojima et al. (1998)
reported that DEHP did not activate either FATP1 or FABPpm in mouse liver, this current investigation demonstrated that FATP1 was significantly induced by DEHP, MEHP, and EHA by 1.7- to 2.5-fold. Many factors might contribute to the inconsistency of induction response (e.g., specifies difference, cell line/tissue utilized, experimental conditions), which reflects the complexity of the activation mechanism.
HFABP is a member of the cytoplasmic fatty acid binding protein family, a large family containing small cytoplasmic proteins responsible for the intracellular trafficking of fatty acids, and is believed to play an integral role in the metabolism, transport, and membrane incorporation of fatty acids (Knipp et al., 1999; Soares et al., 1987
). It was suggested that HFABP might participate in the placental transfer of fatty acids to the fetus due to its specific distribution to the labyrinthine zone of the rat placenta (Knipp et al., 2000
). The expression levels of the HFABP mRNA and protein were up-regulated in a dose- and time-dependent manner when exposed to DEHP, MEHP, and EHA. It should be noted that the maximum fold change in levels observed for HFABP were lower and occurred at a later time when compared to FATP1. These differences might suggest differential fatty acid transport regulation and/or the possible involvement of other unidentified regulators, e.g., PPAR
co-activator 1 (PGC-1) (Storey, 2003
). Interestingly, the consistent effects of DEHP and its metabolites on FATP1 and HFABP did suggest coordinated regulation of their expression, which has been shown to occur through PPAR
transcriptional regulation.
Notably, induction of the mRNA of the majority of these target genes in this study reached peak levels at 2 or 4 h and then diminished to the levels below those of the control cells at 24 h. The highest protein expression levels were observed at 12 or 24 h, consistent with the nature of post-transcriptional events. While this has not been extensively investigated here, we did see some qualitative indications of apoptosis at 12 or 24 h in our study, including shrinkage of the cell membrane and blebbing on the membrane surface under the microscope (data not shown). This may explain the reduction in the mRNA levels at the 24-h time period. It has been previously reported that DEHP and MEHP can cause rat germ cell apoptosis through Fas and other death-associated receptors (Giammona et al., 2002; Richburg et al., 2000
). Further experiments like DNA fragmentation studies may be utilized to confirm these findings. Recently, Yokoyama (Yokoyama et al., 2003
) further demonstrated that the apoptosis induced by MEHP in U937 cells was in part caused by PPAR
(not PPAR
) activation. These findings imply that continued exposure to DEHP and its metabolites might cause concurrent physiological changes through alternate signal transduction pathways.
The up-regulation of FATP1 and HFABP, known mediators of active LCFA uptake/transport, consisted with the significantly increased uptake of polyunsaturated EFAs (AA, DHA, LA, ALA) when compared with saturated SA and monounsaturated OA in the DEHP-, MEHP-, and EHA-treated HRP-1 cells (Fig. 6). The fact that Pmono,influx and Pmono,efflux of AA and DHA were both increased under exposure to DEHP and its metabolites indicates the up-regulation of the fatty acid transport in both the apical and basolateral membranes of HRP-1 cells. This is, again, in accordance with the membrane localization of FATP1, which has been demonstrated to be expressed in the BBM (brush-border membrane) and BPM (basal-plasma membrane) of the human placental cells (Dutta-Roy, 2000; Haggarty, 2002
). The influx ratio of the representative
-3 (i.e., DHA) and
-6 (i.e., AA) EFA were differentially induced upon treatment (Fig. 7). This differential effect of DEHP, MEHP, and EHA on
-3 and
-6 EFA uptake/transport in placental trophoblastic model might be due to the up-regulation of the PPARs and the subsequent change in PPAR-regulated fatty acid transporters. Other mechanisms including the potential regulation effects of DEHP and its metabolites on placental EFA metabolism enzymes via PPAR mediation might play a role (Bocher et al., 2002
; Lemberger et al., 1996
).
Overall, the uptake/transport results imply that EFA nutrition supply to the fetus via the placenta might be altered upon maternal exposure to DEHP and its metabolites, and these effects might differ with respect to the degree of saturation and the classification of EFAs. These results may suggest a potential for altering fetal development, since different EFAs have long been suggested to have differential effects on early neonate development (Bocher et al., 2002; Innis, 2003
; Uauy et al., 1999
). For example, DHA is critical for myelin synthesis and is linked to cognition and other neurological disorders (Innis, 2003
; Uauy et al., 1999
). Furthermore, the maternal fatty acid/lipid homeostasis environment might have dramatic changes upon exposure to phthalates at this period (Uauy et al., 1999
) (e.g., composition and distribution of maternal fatty acids in the blood and the adipose tissue).
In summary, this study demonstrated that the environmental contaminant DEHP and/or its metabolites could alter the expression of PPAR and PPAR
and major fatty acid transferring proteins (FATP1, HFABP), subsequently increasing the uptake/transport of LCFAs in HRP-1 rat trophoblastic cells. Subsequent changes in placental fatty acid homeostasis may lead to adverse fetal development (i.e., birth defects and potentially fetal mortality). Our work provides a valuable model for further investigating the functional and physiological significance of the effects of xenobiotics on EFA homeostasis and proper fetal development, which may eventually provide prevention/correction methods of abnormal fetal EFA-related congenital defects.
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ACKNOWLEDGMENTS |
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