Differential effects of low-dose docosahexaenoic acid and eicosapentaenoic acid on the regulation of mitogenic signaling pathways in mesangial cells

Differential effects of low-dose docosahexaenoic acid and eicosapentaenoic acid on the regulation of mitogenic signaling pathways in mesangial cells AHAD N. K. YUSUFI, JINGFEI CHENG, MICHAEL A. THOMPSON, HENRY J. WALKER, CATHERINE E. GRAY, GINA M. WARNER, and JOSEPH P. GRANDE ROCHESTER, MINNESOTA Although dietary fish oil supplementation has been used to prevent the progression of kidney disease in patients with IgA nephropathy, relatively few studies provide a mechanistic rationale for its use. Using an antithymocyte (ATS) model of mesangial proliferative glomerulonephritis, we recently demonstrated that fish oil inhibits mesangial cell (MC) activation and proliferation, reduces proteinuria, and decreases histologic evidence of glomerular damage. We therefore sought to define potential mechanisms underlying the antiproliferative effect of docosahexaenoic acid (DHA) and eicosapentaenoic acid (EPA), the predominant ␻-3 polyunsaturated fatty acids found in fish oil, in cultured MC. DHA and EPA were administered to MC as bovine serum albumin fatty-acid complexes. Low-dose (10-50 ␮mol/L) DHA, but not EPA, inhibited basal and epidermal growth factor (EGF)–stimulated [3H]-thymidine incorporation in MCs. At higher doses (100 ␮mol/L), EPA and DHA were equally effective in suppressing basal and EGF-stimulated MC mitogenesis. Low-dose DHA, but not EPA, decreased ERK activation by 30% (P < .01), as assessed with Western-blot analysis using phosphospecific antibodies. JNK activity was increased by low-dose DHA but not by EPA. p38 activity was not significantly altered by DHA or EPA. Cyclin E activity, as assessed with a histone H1 kinase assay, was inhibited by low-dose DHA but not by EPA. DHA increased expression of the cell cycle inhibitor p21 but not p27; EPA had no effect on p21 or p27. We propose that the differential effect of low-dose DHA vs EPA in suppressing MC mitogenesis is related to down-regulation of ERK and cyclin E activity and to induction of p21. (J Lab Clin Med 2003;141:318-30) Abbreviations: ATS ⫽ antithymocyte serum; BSA ⫽ bovine serum albumin; DHA ⫽ docosahexaenoic acid; ECL ⫽ enhanced chemiluminescence; EDTA ⫽ ethylenediaminetetraacetic acid; EGF ⫽ epidermal growth factor; EPA ⫽ eicosapentaenoic acid; HEPES ⫽ N-2-hydroxyethylpiperazine-N-2-ethanesulfonic acid; IL-6 ⫽ interleukin-6; ITS⫹ ⫽ insulin, transferrin, selenium, and BSA; LDH ⫽ lactate dehydrogenase; MAPK ⫽ mitogen-activated protein kinases; MC ⫽ mesangial cells; PBS ⫽ phosphate-buffered saline solution; PDGF ⫽ platelet-derived growth factor; PMSF ⫽ phenylmethylsulfonylfluoride; pRb ⫽ retinoblastoma protein; ␻-3 PUFA ⫽ ␻-3 polyunsaturated fatty acid; RIPA ⫽ PBS, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS, 100 ␮g/mL PMSF, 2 ␮g/mL aprotinin, and 200 ␮mol/L sodium orthovanidate; SDSPAGE ⫽ sodium dodecyl sulfate–polyacrylamide gel electrophoresis; TBS ⫽ Tris-buffered saline solution; TCA ⫽ trichloroacetic acid; TdT ⫽ terminal deoxynucleotidyl transferase; TNF ⫽ tumor necrosis factor; VSMC ⫽ vascular smooth muscle cells From the Renal Pathophysiology Laboratory, Department of Laboratory Medicine and Pathology, Mayo Clinic. Supported by National Institutes of Health grants DK16105 and 55603. Submitted for publication August 15, 2002; revision submitted December 1, 2002; accepted December 9, 2002. 318 Reprint requests: Joseph P. Grande, MD, PhD, Mayo Clinic and Foundation, 200 First Street SW, Rochester, MN 55905; e-mail: grande.joseph@mayo.edu. Copyright © 2003 by Mosby, Inc. All rights reserved. 0022-2143/2003/$30.00 ⫹ 0 doi:10.1016/S0022-2143(03)00005-2 J Lab Clin Med Volume 141, Number 5 Recent studies have demonstrated that dietary supplementation with ␻-3 PUFA retards disease progression in human and experimental renal disease.2–12 Fish and marine oils, including EPA (C20:5␻3) and DHA (C22: 6␻3), are abundant sources of ␻3 PUFAs.13,14 Fish oil has been shown to reduce blood pressure, reduce serum lipid levels, decrease eicosanoid and cytokine production, and reduce proteinuria in human and experimental models of renal disease.9,15–22 In IgA nephropathy, the most common glomerulonephritis worldwide,23 the rate of renal disease progression was significantly reduced in patients given a fish oil supplement containing EPA and DHA.10 –12,24 In the ATS model of mesangial proliferative glomerulonephritis, we found that fish oil inhibits mesangial activation and proliferation, reduces proteinuria, and decreases histologic evidence of glomerular damage.1 These studies suggest that fish oil protects against renal disease progression by inhibiting the proliferative response of MC to injury. However, the mechanism by which fish oil inhibits MC proliferation has not been elucidated. In cultured cells, DHA and EPA may inhibit cell growth by decreasing the production of growth factors/ cytokines, by inhibiting mitogenic signaling cascades, or by triggering apoptosis.25–29 In many studies, relatively high doses of fatty acids (⬎50 ␮mol/L) have been used to demonstrate an inhibitory effect of EPA and DHA on mitogenesis.30 –32 Furthermore, the route of administration of fatty acids to cultured cells may significantly affect experimental results. For example, direct administration of fatty acids to culture medium may reduce binding of growth factors to their cognate receptors through a detergent-like effect.33 To avoid this complication, we administered DHA and EPA to MC as a BSA–fatty acid complex. In our previous study, we found that complexes containing low doses (10-20 ␮mol/L) of EPA and DHA were readily incorporated into MC plasma membranes and tended to replace arachidonic acid as a membrane constituent. Although both DHA and EPA were incorporated into MC plasma membranes, we found that only low-dose DHA (10-20 ␮mol/L) inhibited basal and PDGF-stimulated mitogenesis of MCs; an equimolar dose of EPA was without effect.1 The basis for this differential effect of low-dose DHA and EPA on MC mitogenesis has not been previously established. The main objective of this study was to identify potential sites in mitogenic signaling pathways that are differentially regulated by low-dose (20 ␮mol/L) DHA and EPA. This information is essential to providing the basis for studies to establish the mechanism whereby fish oil suppresses MC mitogenesis. We demonstrate that the antiproliferative effect of DHA is associated with down-regulation of ERK, inhibition of cyclin E- Yusufi et al 319 cdk2 activity, and up-regulation of the cell cycle inhibitor p21. The antiproliferative effect of DHA is not associated with apoptosis in MC. In accordance with observations made by others,31 at higher doses, we found that both DHA and EPA inhibit MC mitogenesis. We propose that the protective effect of fish oil in preventing disease progression in IgA nephropathy and other mesangial proliferative renal diseases is at least in part the result of a suppressive effect of DHA on MC mitogenesis. METHODS [3H]-thymidine was purchased from Du Pont/ New England Nuclear Research Products (Boston, Mass). Primary antibodies (eg, for p-ERK, p-p38, p21, p27, cyclin D1, cyclin E, cdk-2, cdk-4) and horseradish-peroxidase-conjugated secondary antibodies were obtained from Santa Cruz Biotechnology, Inc (Santa Cruz, Calif). The ␤-actin antibody was obtained from Sigma Chemical Co (St Louis, Mo). EPA and DHA were from Cayman Chemical (Ann Arbor, Mich). Protein A agarose was obtained from Santa Cruz Biotechnology. Histone H1 was obtained from Calbiochem (La Jolla, Calif). Other reagents and supplies were obtained through standard commercial suppliers. Preparation of fatty acid–albumin complexes. Fatty acid–BSA complexes were prepared as previously described.1 In brief, fatty acids were resuspended in absolute ethanol and slowly added to a 0.3 mmol/L solution of essential fatty acid–free BSA (Sigma) in PBS, which was stirred under liquid nitrogen for 5 hours. The final molar ratio of fatty acid to albumin was approximately 0.7:1.0. Solutions were aliquotted, stored at ⫺80°C, and thawed immediately before being added to MC cultures. MC culture. MC cultures were obtained from 200 g male Sprague-Dawley rats by means of differential sieving, as previously described.34 –36 This protocol was approved by the Institutional Animal Welfare Committee of the Mayo Clinic and Foundation in accordance with the principles of laboratory animal care (NIH publication no. 86-23, revised 1992). In brief, rats were anesthetized with an intraperitoneal injection of a 1:1 mixture of 20 mg/mL xylazine and 100 mg/mL ketamine. The kidneys were excised, the renal capsule removed, and the cortical tissue minced and passed through a stainless-steel sieve (200 ␮m pore size). The homogenate was sequentially sieved through nylon meshes with 390, 250, and 211 ␮m pore openings. We then passed the cortical suspension over a 60 ␮m sieve to collect glomeruli. Putative glomerular preparations were evaluated with the use of light microscopy. Preparations typically contained more than 90% glomeruli. Glomeruli were seeded on plastic tissue-culture dishes and grown in complete Waymouth’s medium (Waymouth’s medium supplemented with 20% heat inactivated fetal calf serum, 15 mmol/L HEPES, 1 mmol/L sodium pyruvate, 0.1 mmol/L nonessential amino acids, 2 mmol/L L-glutamine, 100 IU/mL penicillin, 100 ␮g/mL streptomycin, and 1% ITS⫹). Fresh medium was added every 3 days. Cell outgrowths were characterized as MC on the basis of positive Materials. 320 Yusufi et al J Lab Clin Med May 2003 Fig 1. Low dose of DHA, but not EPA, suppresses basal and EGF-stimulated mitogenesis of MC. MC were treated with 10, 50, or 100 ␮mol/L BSA (hatched bars), DHA (black bars), or EPA (white bars) for 24 hours in the absence (A) or presence (B) of EGF (20 ng/mL) before assessment of [3H]-thymidine uptake. Data expressed as mean ⫾ SEM (n ⫽ 3 experiments, each performed in duplicate). *Significantly different from BSA-treated control (A) or EGF-stimulated BSA control (B) (P ⬍ .05). immunohistochemical staining for vimentin and smooth muscle–specific actin, along with negative staining for cytokeratin, factor VIII–related antigen, and leukocyte-common antigen (antibodies from Dako Corp, Carpinteria, Calif). MCs were passed once a week after treatment with trypsin-EDTA (0.02%). Cells used in experiments were from passages 5 through 12. [3H]-thymidine incorporation. MC were plated into 24well tissue-culture dishes, 5 ⫻ 104 cells/well and grown for 24 to 48 hours in complete Waymouth’s medium. Cells were re-fed with Waymouth’s medium containing 0.5% calf serum and supplemented with fatty acids (EPA or DHA, 10-100 ␮mol/L). The fatty acid–BSA conjugates are rapidly taken up by MCs and incorporated into membrane phospholipids.1 Control cultures were given equimolar concentrations of lipid-free BSA. After 20 hours, cells were treated with methyl[3H]-thymidine (1 ␮Ci/mL), after which cultures were incubated for an additional 4 hours. As indicated, EGF (20 ng/ mL) was added at the time of fatty acid administration. Cells were washed twice with PBS and subjected to lysis by means of addition of 0.2N NaOH. After 20 minutes, the cell lysate was neutralized with HCl. TCA was added to a final concentration of 10%. The solution was passed over glass-fiber disks (GF/C; Whatman, Clifton, NY), which were then washed J Lab Clin Med Volume 141, Number 5 Yusufi et al 321 Fig 2. DHA, but not EPA, inhibits ERK activation. MC were treated with BSA (control), 20 ␮mol/L DHA (black bars), or 20 ␮mol/L EPA (white bars) for 2 and 24 hours. p-ERK was assessed by Western blot with phosphospecific antibody, as described in the Methods. Data expressed as percentage of BSA-treated control (100%) (mean ⫾ SEM; n ⫽ 3 experiments, each performed in duplicate). *Significantly different from BSA-treated control (P ⬍ .01). Inset: blot of a representative experiment. twice with 10% TCA and once with 70% ethanol. We assayed radioactivity on the disks with the use of liquid scintillation counting. Incorporation of [3H]-thymidine was used as a measure of the rate of mitogenic synthesis of DNA. LDH assay. We assessed cell viability after incubation with fatty acid–BSA complexes with the use of a LDH assay (procedure 228-UV; Sigma Diagnostics, St Louis, Mo). LDH activity was calculated from the change in absorbance at 340 nm/min. Western-blot analysis. MC cultures were treated with fatty acid–albumin complexes (20 ␮mol/L for 2 or 24 hours) as described above. After incubation, MCs were rinsed, harvested, and subjected to sonication (three cycles of 10 seconds each, 8 ␮m amplitude) in RIPA homogenizing buffer. The homogenates were centrifuged at 11,000g for 20 minutes. Protein concentration was determined with the method of Lowry et al.37 Equal amounts of lysate proteins (30 ␮g) were subjected to SDS-PAGE in the PROTEAN II minigel system (BioRad, Hercules, Calif). Lysates were denatured for 3 minutes at 95°C in SDS loading buffer in accordance with the method of Laemmli et al.38 Electrophoresis was performed at a constant current (200 mA/gel) and followed by transfer to nitrocellulose membranes. The membranes were blocked with 5% nonfat dry milk in TBS containing 0.5% Tween 20, followed by incubation with appropriate primary antibodies and horseradish-peroxidase-conjugated secondary antibodies. We then visualized the blots by exposing them to x-ray film using an ECL kit (Amersham-Pharmacia Biotech, Inc, Piscataway, NJ). Transfection studies. We measured JNK activity with a transfection-based in vivo kinase assay kit (Clonetech Laboratories, Inc, Palo Alto, Calif), in accordance with the manufacturer’s instructions. In brief, MC were plated into 24-well culture dishes at 8 ⫻ 104 cells/well in complete Waymouth’s medium. Twenty-four hours after plating, cells were cotransfected with a transactivator expression vector (pTet-JUN), a firefly luciferase reporter vector (pTRE-Luc), and a control Renilla luciferase reporter vector. We performed transfections with FuGENE 6 Transfection Reagent (Roche Molecular Biochemical, Indianapolis, Ind). Eighteen hours after transfection, BSA-conjugated DHA or EPA (20 ␮mol/L) was added. Control cells received BSA only. Cells were rinsed and subjected to lysis after 2 and 24 hours’ treatment. We assessed luciferase activity with the Dual-Luciferase Reporter Assay System (Promega Corp, Madison, Wis). In vitro kinase assays. The p44/42 MAP Kinase Assay Kit (Cell Signaling Technology, Inc, Beverly, Mass) was used to measure ERK kinase activity, in accordance with the 322 Yusufi et al J Lab Clin Med May 2003 Fig 3. DHA, but not EPA, stimulates JNK activation. MC were treated with BSA (control), 20 ␮mol/L DHA (black bars), or 20 ␮mol/L EPA (white bars) for 2 and 24 hours. JNK activity was assessed by a transfectionbased in vivo kinase assay, as described in the Methods. Data expressed as percentage of BSA-treated control (100%) (mean ⫾ SEM; n ⫽ 3 experiments, each performed in duplicate). *Significantly different from BSA-treated control (P ⬍ .01). manufacturer’s instructions. In brief, after treatment with DHA or EPA (20 ␮mol/L; 2 and 24 hours), MC were rinsed, harvested, and sonicated four times, 5 seconds each time, in 1⫻ lysis buffer plus 1 mmol/L PMSF. Samples were microcentrifuged for 10 minutes at 4°C, and protein concentrations in the supernatants were determined as described above. Cell lysate (200 ␮L) containing 200 ␮g total protein was added to 15 ␮L of resuspended immobilized phospho-p44/42 MAP kinase (Thr202/Tyr204) monoclonal antibody and incubated with gentle rocking overnight at 4°C. After samples were microcentrifuged for 30 seconds at 4°C, pellets were washed twice with 1⫻ lysis buffer and twice with 1⫻ kinase buffer. The washed pellets were suspended in 50 ␮L 1⫻ kinase buffer supplemented with 200 ␮mol/L ATP and 2 ␮g Elk-1 fusion protein, then incubated for 30 minutes at 30°C. Reactions were terminated with 25 ␮L of 3⫻ SDS sample buffer. Samples were boiled for 5 minutes, vortexed, microcentrifuged for 2 minutes, and then loaded (30 ␮L) on SDS-PAGE gels (12%). We analyzed samples with the use of Western blotting, as described above. Histone H1 kinase assays for cyclin-cdk activity. Fatty acid–treated MC (20 ␮mol/L; 2 and 24 hours) were rinsed and subjected to lysis in RIPA buffer, after which protein concentrations were determined, as described above. Equal amounts of lysate protein (200 ␮g) were immunoprecipitated with antibodies specific for cyclin D1 and cyclin E. The immune complexes were collected with protein A–agarose and washed twice with RIPA buffer. Complexes were resuspended and washed twice with kinase buffer (50 mmol/L Tris-HCl [pH 7.4], 10 mmol/L MgCl2, 1 mmol/L dithiothreitol). Complexes were then resuspended in 50 ␮L kinase buffer containing 2 ␮g histone H1, 200 ␮mol/L ATP, and 5 ␮Ci [␥-32P]ATP (3000 Ci/mmol) and incubated at 30°C for 30 minutes. After incubation, 25 ␮L of 3⫻ SDS loading buffer was added and the samples were boiled and subjected to electrophoresis on a 12% SDS-PAGE gel. The gels were dried, after which incorporation of 32P was visualized with the use of autoradiography and quantitated with a Kodak image analysis system (Eastman Kodak Co, Rochester, NY). Assays for apoptosis. Structural changes in the nuclear chromatin of fatty acid–treated MCs (20-100 ␮mol/L, 48 hours) undergoing apoptosis were detected on staining with bisbenzimide (Hoechst 33342; Calbiochem). Cells were pelleted at 300g, washed with PBS, and fixed in 1% gluteraldehyde in PBS for 30 minutes at room temperature. Cells were then aliquoted onto glass slides, stained with Hoechst 33342 (10 ␮g/mL in deionized water) at room temperature for 30 minutes, and rinsed with PBS. Slides were coverslipped with Permafluor mounting medium (Thermo Shandon, Pittsburgh, Penn) and analyzed with an Olympus fluorescence microscope (Olympus, Melville, NY). Cells containing three or more chromatin fragments were considered apoptotic. Apoptosis was assessed with the ApoTag⫹ peroxidase in situ apoptosis detection kit (Intergen, Purchase, NY). In brief, MCs were pelleted at 300g and washed with PBS. ApoTag assay–treated and control cells were aliquoted onto poly-Llysine– coated glass slides and fixed in 1% methanol-free formaldehyde at room temperature for 30 minutes. Cells were then postfixed in ethanol/acetic acid (2:1), washed, and treated with 3% H2O2 to quench endogenous fluorescence. After rinsing in PBS, equilibration buffer was applied for 3 minutes before the addition of TdT. The reaction was developed with diaminobenzidine and counterstained with methyl green. For negative control slides, we omitted the TdT enzyme step. Caspase-3 activity in control and fatty acid–treated (20-100 J Lab Clin Med Volume 141, Number 5 Yusufi et al 323 Fig 4. Neither DHA nor EPA alters p38 activation. MC were treated with BSA (control), 20 ␮mol/L DHA (black bars), or 20 ␮mol/L EPA (white bars) for 2 and 24 hours. p-p38 was assessed by Western blot with phospho specific antibody, as described in the Methods. Data expressed as percentage of BSA-treated control (100%) (mean ⫾ SEM; n ⫽ 3 experiments, each performed in duplicate). *Significantly different from BSA-treated control (P ⬍ .05). Inset: blot of a representative experiment. ␮mol/L, 24 hours) MC was determined fluorometrically with the CaspACE Assay System (Promega). Statistical analysis. The data presented herein are representative of at least three independent experiments. Statistical analysis was performed with the use of In Stat (Graph Pad, San Diego, Calif). Pairwise comparisons between DHA- or EPA-treated and control cells were evaluated with the use of Student’s t test. P values of less than .05 were considered statistically significant. Results Inhibitory effect of DHA and EPA on MC mitogenesis is dose-dependent. The dose-dependent effect of DHA and EPA on basal and EGF-stimulated MC mitogenesis was assessed on the basis of [3H]-thymidine incorporation, as described in the Methods. In accord with our previous observations,1 10 ␮mol/L DHA significantly inhibited basal and EGF-stimulated MC mitogenesis (⫺33% and ⫺41% respectively; P ⬍ .05), whereas EPA was without effect (Fig 1). At a dose of 100 ␮mol/L, both DHA and EPA, administered as BSA– fatty acid conjugates, inhibited MC proliferation to a similar extent. BSA alone (10, 50, or 100 ␮mol/L) had no significant effect on basal or EGF-stimulated MC mitogenesis. The antiproliferative effect of DHA was not a result of cytotoxicity; no appreciable release of LDH into culture supernatant from cells treated with DHA, EPA, or BSA was observed (data not shown). DHA inhibits ERK activation but stimulates the JNK path- We assessed the effect of 20 ␮mol/L DHA or EPA on p-ERK and p-p38 expression by conducting Western blotting of cell lysates with phosphospecific antibodies. JNK activity was assessed with a transfection-based in vivo kinase assay, as described in the Methods. The antiproliferative effect of 20 ␮mol/L DHA was associated with a significant decline in pERK expression (⫺30% after 2 hours’ incubation, ⫺29% after 24 hours; P ⬍ .01). In contrast, 20 ␮mol/L EPA, which had no effect on MC mitogenesis, had no significant effect on p-ERK expression (Fig 2). These findings were confirmed with an in vitro kinase assay for ERK activity; 24 hours’ treatment with DHA, but not EPA, inhibited ERK activation (data not shown, P ⬍ .05). Treatment of MC with 20 ␮mol/L DHA significantly way. 324 J Lab Clin Med May 2003 Yusufi et al Fig 5. DHA, but not EPA, inhibits cyclin E kinase activity. MC were treated with BSA (control), 20 ␮mol-L DHA (black bars), or 20 ␮mol/L EPA (white bars) for 2 and 24 hours. Cyclin E kinase activity (A) was assessed by histone H1 kinase assay; cyclin E levels (B) were assessed by Western blot with phosphospecific antibody, as described in the Methods section. Data expressed as percentage of BSA-treated control (100%) (mean ⫾ SEM; n ⫽ 2-3 experiments, each performed in duplicate). *Significantly different from BSA-treated control (P ⬍ .01). Insets: blots of representative experiments. induced JNK activity (40% after 2 hours’ incubation; P ⬍ .01). Treatment of MC with 20 ␮mol/L EPA had no significant effect on JNK activity (Fig 3). Neither EPA nor DHA significantly affected p-p38 expression (Fig 4). Effect of DHA and EPA on cell cycle–regulatory proteins. Progression of the cell cycle from G1 to S-phase is mediated through activation of cyclin D-cdk4,6 and cyclin E-cdk2 complexes. We assessed the role of DHA and EPA on the activity of cyclin D and cyclin E with Western blotting and the histone H1 kinase assay. DHA significantly suppressed cyclin E kinase activity (28% after 2 hours; P ⬍ .01). Cyclin E levels and cyclin E kinase activity were not altered by 20 ␮mol/L EPA (Fig 5). cdk2 levels did not significantly change after treatment with DHA or EPA (data not shown). Neither DHA nor EPA had a significant effect on cyclin D kinase activity, cyclin D levels (Fig 6), or cdk4 levels (data not shown). Effect of DHA and EPA on the cell cycle–inhibitory proteins p21 and p27. The antiproliferative effects of 20 ␮mol/L DHA were associated with induction of the cell-cycle inhibitor p21 (51% after 6 hours’ incubation, 73% after 24 hours; P ⬍ .01). Treatment of MC with 20 ␮mol/L EPA, which had no effect on MC proliferation, likewise had no effect on p21 levels (Fig 7). Expression of p27 was not significantly altered by treatment with DHA or EPA (Fig 8). DHA and EPA do not induce apoptosis in MC. Apoptosis was assessed in MC treated with 20 to 100 ␮mol/L DHA or EPA by means of Hoechst 33342 staining, ApoTag⫹ assay, and CaspACE 3 assay, as described in the Methods. Under these experimental conditions, we detected no structural evidence of apoptosis (chromatin condensation and fragmentation, TUNEL positivity). Caspase-3 activity in treated cells did not differ significantly from that in BSA-treated controls (three independent experiments, data not shown). As a positive control, we treated MC with 20 ng/mL TNF-␣ and 10 ␮g/mL cycloheximide for 24 hours; these showed extensive chromatin condensation and nuclear fragmentation. Caspase-3 activity in the positive control cells was significantly increased (95%; data not shown). DISCUSSION Previous clinical and experimental studies have provided evidence that fish oil has a role in the treatment of IgA nephropathy and other progressive renal diseas- J Lab Clin Med Volume 141, Number 5 Yusufi et al 325 Fig 6. Neither DHA nor EPA alters cyclin D kinase activity or cyclin D levels. MC were treated with BSA (control), 20 ␮mol/L DHA (black bars), or 20 ␮mol/L EPA (white bars) for 2 and 24 hours. Cyclin D kinase activity (A) was assessed by histone H1 kinase assay; cyclin D levels (B) were assessed by Western blot, as described in the Methods section. Values are expressed as percentage of BSA-treated control (100%) and represent the mean ⫾ SEM (n ⫽ 2-3 experiments, each performed in duplicate). Insets: blots of representative experiments. es.3,11,12 However, the mechanisms underlying the protective effect of fish oil have not been established. We have previously demonstrated that low doses of DHA and EPA, the predominant long-chain PUFA in fish oil, are readily incorporated into MC membranes.1 When administered as fatty acid–BSA conjugates, DHA and EPA tend to replace arachidonic acid as membrane phospholipids in MC. At doses of 10 to 20 ␮mol/L, DHA is a potent inhibitor of MC mitogenesis, whereas EPA is without effect. The inhibitory effect of DHA on MC proliferation is not a result of cytotoxicity, as assessed on the basis of LDH release. The antiproliferative effect of DHA is likely the result of modulation of mitogenic signaling pathways. These potential targets of DHA have not been previously defined in MC. The main objective of our study was to determine which mitogenic signaling pathways and cell cycleregulatory proteins are differentially regulated by lowdose (20 ␮mol/L) DHA or EPA. In our initial studies, we characterized in more detail the dose-dependence of the inhibitory effects of DHA and EPA on basal and EGF-stimulated MC mitogenesis. We found that low doses of DHA (10-20 ␮mol/L) are effective in inhibiting MC mitogenesis, whereas equimolar concentrations of EPA are without effect. At higher doses (100 ␮mol/L), both DHA and EPA were equally effective in inhibiting MC mitogenesis. These observations are in agreement with those of other investigators, who have shown that higher doses of EPA (50-100 ␮mol/L) are effective in inhibiting proliferation of endothelial cells,39 VSMC,40,41 and MC.31 Our subsequent studies, designed to define the role of DHA in mitogenic signaling pathways, were conducted with doses of 20 ␮mol/L, at which a differential effect of DHA and EPA on mitogenesis was observed. We studied the role of DHA and EPA in MAPK pathways. MAPK are key regulators of cell growth and apoptosis; they include ERK, p38, and JNK.42,43 To account for the possibility that DHA or EPA transiently modulates MAPK signaling pathways, we chose to analyze the effects of the ␻-3 PUFA on MAPK activity 2 and 24 hours after treatment. We selected the 2-hour time point to allow time for MC cultures to take up the fatty acid–BSA complexes and incorporate them into plasma membrane phospholipids. MAPK activity was also assessed after 24 hours of treatment, the time at which [3H]-thymidine-uptake studies were performed. We found that the antiproliferative effect of DHA was associated with significant inhibition of ERK activity, as assessed with two complementary methods: immunoblotting with a phospho-ERK antibody and an in vitro kinase assay. We observed the inhibitory effect of DHA on ERK activity after both 2 and 24 hours of treatment. An equimolar dose of EPA, which did not 326 Yusufi et al J Lab Clin Med May 2003 Fig 7. DHA, but not EPA, induces expression of the cell-cycle inhibitor p21. MC were treated with BSA (control), 20 ␮mol/L DHA (black circles), or 20 ␮mol/L EPA (white triangles) for 2, 6, and 24 hours. p21 expression was assessed by Western blot, as described in the Methods section. Data expressed as percentage of BSA-treated control (100%) (mean ⫾ SEM; n ⫽ 3 experiments, each performed in duplicate). *Significantly different from BSA-treated control (P ⬍ .01). Inset: blot of a representative experiment. inhibit mitogenesis, had no effect on ERK activity. ERK activation is recognized as a critical mitogenic signaling pathway that directs growth of cells in response to a wide variety of mitogens, including PDGF and EGF. Whereas ERK activation is characteristically associated with growth-signaling pathways, the activation of p38 and JNK has been associated with apoptosis.44 Previous studies have indicated that differential activation of the ERK versus JNK or p38 pathways may determine whether a cell will proliferate or undergo apoptosis.45 For example, in PC12 cells, concurrent activation of JNK and p38 kinase pathways and inhibition of the ERK pathway induce apoptosis, whereas direct and selective activation of the ERK pathway prevents apoptosis.44 Activation of ERK may prevent apoptosis in response to JNK activation.46 We found that DHA inhibited ERK and transiently activated the JNK pathway without triggering MC apoptosis. Other investigators have shown that prolonged activation of JNK is necessary to trigger apoptosis in MC.47 The effects of DHA or EPA on apoptosis appear to be cell type–specific. For example, in VSMC, DHA induces apoptosis by way of activation of p38.48 Higher doses (40-80 ␮mol/L) of DHA in VSMC induce caspase 3 activity and promote nuclear condensation, a structural feature of apoptosis.49 DHA also induces apoptosis in Jurkat leukemia T-cells and colon cancer cells.50 DHA and EPA may promote apoptosis of tumor cells by way of lipid peroxidation.29 However, DHA inhibits sphingosine-induced apoptosis in HL60 cells.51 DHA inhibits TNF-␣–induced apoptosis of human monocytic U937 cells52 and neuronal cells.53 Notably, EPA does not inhibit apoptosis of HL60 cells, indicating that DHA and EPA differentially regulate apoptosis.51 The effects of DHA and EPA on MC apoptosis have not been described previously. J Lab Clin Med Volume 141, Number 5 Yusufi et al 327 Fig 8. Neither DHA nor EPA alters expression of the cell-cycle inhibitor p27. MC were treated with BSA (control), 20 ␮mol/L DHA (black circles), or 20 ␮mol/L EPA (white triangles) for 2, 6, and 24 hours. p27 expression was assessed by Western blot, as described in the Methods section. Data expressed as percentage of BSA-treated control (100%) (mean ⫾ SEM; n ⫽ 3 experiments, each performed in duplicate). Inset: blot of a representative experiment. In mammalian cells, cell cycle progression is regulated through sequential activation of cyclin-cyclin– dependent kinase complexes.54 –56 A major point of regulation of the cell cycle is in the G1-to-S transition. When a cell is stimulated to proliferate, cyclin D associates with the cyclin-dependent kinases cdk4 and cdk6, whereas cyclin E associates with cdk2. Both cyclin D– cdk4/cdk6 and cyclin E– cdk2 phosphorylate the pRb.57,58 Activation of both cyclin D and cyclin E is essential for progression from G1 to S-phase of the cell cycle.59,60 We found that DHA transiently reduced cyclin E activity but had no significant effect on cyclin D activity. No significant changes in cyclin D, cyclin E, cdk2, or cdk4 levels were observed after DHA or EPA treatment. In melanoma cells, DHA promotes cell-cycle arrest and apoptosis in association with decreased pRb phosphorylation.61 In HT-29 colon cancer cells, DHA inhibits proliferation by preventing activation of both cyclin D-cdk and cyclin E– cdk complexes.32 In VSMC, high-dose (80-160 ␮mol/L) EPA and DHA inhibit proliferation by inhibiting phosphorylation of the cyclin E– cdk2 complex.30 On the basis of our findings, we conclude that at low doses (10-20 ␮mol/ L), the differential effect of DHA versus that of EPA on MC mitogenesis is related to inhibition of cyclin E– cdk2 activity by DHA but not by EPA. Activity of cyclin– cdk complexes is regulated by two families of cdk-inhibitory proteins: the inhibitors of cdk (INK) family, which includes p15, p16, p18, and p19; and the cdk inhibitory protein (KIP) family, which includes p21, p27, and p57.62 The INK family of cdk inhibitors preferentially binds cdk4 or cdk6, whereas the KIP family blocks the activity of a variety of cyclin-cdk complexes, including cyclin E– cdk2.63 We found that DHA supplementation increased p21 levels. Increased p21 levels were first seen after 6 hours of treatment and remained high after 24 hours. EPA had no significant effect on p21 levels. Neither DHA nor EPA altered p27 levels. Potential mechanisms whereby 328 J Lab Clin Med May 2003 Yusufi et al low-dose DHA or EPA differentially regulate p21 and p27 levels await elucidation. In summary, we demonstrate that low-dose (10-20 ␮mol/L) administration of DHA and EPA differentially modulates MC proliferation. The antiproliferative effect of DHA is associated with down-regulation of ERK and up-regulation of JNK. At these doses, we found no evidence of caspase activation in DHAtreated cells, indicating that the antiproliferative effects are not the result of induction of apoptosis. Cyclin E kinase activity is decreased by DHA but not by EPA. Cell cycle inhibition is associated with significant induction of p21 but not p27. Further studies are needed to determine the mechanism whereby DHA interacts with these critical targets of cell cycle regulation, leading to inhibition of MC proliferation. DHA-mediated “negative crosstalk” with signaling pathways that target the MAPK pathways, cyclin E, p21, or all three may underlie the protective effect of dietary fish oil supplementation in the treatment of chronic glomerular diseases characterized by excessive MC proliferation, such as IgA nephropathy. 10. 11. 12. 13. 14. 15. 16. 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