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 (C20:53) and DHA (C22:
63), 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
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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
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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
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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
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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.
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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-
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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
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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.
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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.
We gratefully acknowledge the excellent secretarial assistance of
Ms. Cherish Grabau.
17.
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