n-6 and n-3 polyunsaturated fatty acids
differentially modulate oncogenic Ras activation in
colonocytes
Esther D.
Collett,
Laurie A.
Davidson,
Yang-Yi
Fan,
Joanne R.
Lupton, and
Robert S.
Chapkin
Molecular and Cell Biology Group, Faculty of Nutrition, and Center
for Environmental and Rural Health, Texas A&M University, College
Station, Texas 77843-2471
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ABSTRACT |
Ras proteins are
critical regulators of cell function, including growth,
differentiation, and apoptosis, with membrane localization of
the protein being a prerequisite for malignant transformation. We have
recently demonstrated that feeding fish oil, compared with corn oil,
decreases colonic Ras membrane localization and reduces tumor formation
in rats injected with a colon carcinogen. Because the biological
activity of Ras is regulated by posttranslational lipid attachment and
its interaction with stimulatory lipids, we investigated whether
docosahexaenoic acid (DHA), found in fish oil, compared with linoleic
acid (LA), found in corn oil, alters Ras posttranslational processing,
activation, and effector protein function in young adult mouse colon
cells overexpressing H-ras (YAMC-ras). We
show here that the major n-3 polyunsaturated fatty acid (PUFA)
constituent of fish oil, DHA, compared with LA (an n-6 PUFA), reduces
Ras localization to the plasma membrane without affecting
posttranslational lipidation and lowers GTP binding and downstream
p42/44ERK-dependent signaling. In view of the central role
of oncogenic Ras in the development of colon cancer, the finding that
n-3 and n-6 PUFA differentially modulate Ras activation may partly
explain why dietary fish oil protects against colon cancer development.
docosahexaenoic acid; linoleic acid; colon cancer; fish oil
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INTRODUCTION |
COLORECTAL CANCER
continues to pose a serious health problem in the United States,
accounting for an estimated 129,400 new cases diagnosed this year, with
~155 deaths each day (34). Over a lifetime, a
person has 1 chance in 18 of developing invasive colorectal cancer
(34). Colorectal cancer evolves from a multistep process
(20) and is a disease strongly influenced by environmental factors, with diet being one of the most important modifying agents (43). Among dietary factors, there are strong
epidemiological, clinical, and experimental data indicating a
protective effect of fish oil-derived n-3 polyunsaturated fatty acids
(PUFA) on colon cancer (2, 10, 35, 43). In general,
dietary lipids rich in linoleic acid (LA; 18:2n-6), found in a variety
of vegetable oils, enhance the development of colon tumors (8,
17, 46, 47), whereas n-3 PUFA-enriched diets, containing
docosahexaenoic acid (DHA; 22:6n-3), reduce colon cancer incidence
(10, 17, 43, 45, 48). This protective effect is exerted at
both the initiation and postinitiation stages of carcinogenesis
(11, 29, 45). Recently, we have shown that the balance
between colonic epithelial cell proliferation and apoptosis can
be favorably modulated by dietary n-3 PUFA, conferring resistance to
toxic carcinogenic agents (11, 29). However, the
underlying molecular mechanism(s) by which distinct classes of dietary
PUFA (n-6 vs. n-3) exert their effects is not known.
Ras genes code for 21-kDa guanine nucleotide binding proteins that play
an important role in colonic epithelial cell growth, differentiation,
and tumor formation (38, 59). Ras proteins (H-Ras, K-Ras
and N-Ras) differ only in the carboxy-terminal region of the
protein, bind guanine nucleotides (GTP and GDP) with high affinity, and
possess intrinsic GTPase activity. Biological Ras activity is
determined by the bound nucleotide (38, 51), which in turn
is regulated by the membrane lipid microenvironment, specifically posttranslational lipid attachment (isoprenylation and palmitoylation) (23, 25) and the interaction with stimulatory lipids
(53). Ras proteins are synthesized in the cytosol on
ribosomes and have a half-life of 24 h (13). After
synthesis, Ras proceeds through a series of posttranslational
processing steps at the COOH terminus that increase protein affinity
for the plasma membrane. Specifically, the H-Ras and N-Ras isoforms are
transported to the plasma membrane through the Golgi, while K-Ras is
transported through the cytosol after cleavage and methylation
(3, 13, 38). The palmitoylation of H-Ras targets the
protein to the plasma membrane in flask-shaped invagination domains
rich in cholesterol and sphingolipids referred to as caveolae
(39, 55). Importantly, we have recently demonstrated that
dietary fish oil reduces colonic Ras membrane localization in
carcinogen-treated rats (16). Because Ras must be
localized to the plasma membrane to be biologically active, and its
constitutive activation directly drives early colonic tumor development
(52, 59), we have determined the action of DHA
(antitumorigenic) vs. LA (protumorigenic) on Ras posttranslational
processing, activation, and signal transduction in colonocytes.
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MATERIALS AND METHODS |
Materials.
RPMI 1640 and Hanks' balanced salt solution were from Mediatech
(Herndon, VA). Fetal bovine serum (FBS) was from Hyclone (Logan, UT).
Insulin/transferrin/selenium (ITS) was obtained from Collaborative Biomedical Products (Bedford, MA). Glutamax and recombinant mouse interferon-
(IFN-
) were from GIBCO-BRL (Grand Island, NY). Fatty acid-free bovine serum albumin (BSA) and 1% penicillin-streptomycin were from Sigma (St. Louis, MO). Fluorescein diacetate and propidium iodide were obtained from Pharmingen (San Diego, CA). Prepoured polyacrylamide gradient gels were from Novex (San Diego, CA). Electroblotting polyvinylidene difluoride (PVDF) membranes were obtained from Millipore (Burlington, MA). Bicinchoninic acid protein assay and SuperSignal chemiluminescent detection reagents were from
Pierce (Rockford, IL). Peroxidase-conjugated secondary antibodies were
from Kirkegaard & Perry (Gaithersburg, MD). Mycoplasma PCR-based ELISA
was purchased from Roche Molecular Biochemical (Indianapolis, IN).
Bromodeoxyuridine (BrdU) labeling kit was from Oncogene Science (Cambridge, MA). The EnzChek caspase 3 assay kit was from Molecular Probes (Eugene, OR). Prepoured silica gel 60 thin-layer chromatography (TLC) plates were from Merck (Darmstadt, Germany). All other reagents were obtained from Sigma.
Cell culture.
Malignant transformed young adult mouse colon (YAMC)-Ras cells
overexpressing v-H-Ras were kindly donated by Dr. Robert Whitehead (Ludwig Institute for Cancer Research, Melbourne, Australia)
(15). Cells (passages 16-20) were
maintained in RPMI 1640 supplemented with 5% FBS, 1% glutamax-1, 1%
ITS, and 1% penicillin-streptomycin at 33°C. The medium was
supplemented with 5,000 U/l of mouse recombinant IFN-
because the
temperature-sensitive mutant SV40 large T antigen gene (tsA58) is under
an IFN-
-inducible promoter, mouse H-2Kb class I gene
(60). For fatty acid treatment, subconfluent cells were
treated for 72 h with 0-100 µM DHA or LA complexed to fatty acid-free BSA (27). Cells treated with BSA plus medium or
medium alone were used as controls for each experiment. All cultures were mycoplasma free, as determined by a PCR-based ELISA.
Cell proliferation.
Cells were plated at an initial density of 20,000 cells per 35-mm dish.
After 24 h, dishes were treated for 72 h with 50 µM DHA or
LA, with media replenished every 24 h. Medium alone was used as a
negative control. After incubation at 33°C, adherent cells were
trypsinized and counted with a hemacytometer. For quantitation of DNA
synthesis, 1-2 × 105 cells were pulsed with BrdU
(1:2,000 dilution, 0.3 µg/well) 3 h before harvest and then
processed according to kit instructions.
Cell viability.
Cells were plated at 20,000 cells per 35-mm dish, and the effect of
fatty acid treatment on viability was assessed with fluorescein diacetate/propidium iodide (31). After incubation with 50 µM DHA or LA for 72 h, cultures were rinsed with
phosphate-buffered saline (PBS) and stained. A minimum of 100 adherent
cells were counted and scored as red (nonviable) or green (viable) with
the use of a Nikon Eclipse TE300 fluorescence inverted microscope equipped with a Princeton Instruments Micromax cooled digital camera
chip and a Metamorph imaging workstation. As a positive control, YAMC
cells (parental cell line) were incubated at a nonpermissive temperature (39°C) overnight. Data were expressed as percent
viability [100 × (no. of green cells/no. of total cells)].
Caspase 3 activity.
Caspase 3 activity was determined with the EnzChek caspase 3 assay kit.
Floating and adherent cells were harvested, and supernatants were
utilized for the activity assay. Fluorescence was measured at a ratio
of 496 to 520 nm 30, 45, 60, and 90 min after substrate addition. Data
were transformed with a log function before statistical analysis. In
addition, a morphological assessment of apoptosis was performed
by acridine orange staining of floating cells as previously described
(5).
Measurement of membrane phospholipid fatty acid composition.
Cell lipids were extracted with chloroform-methanol (2:1 vol/vol), and
individual phospholipid classes were separated by TLC with
chloroform-methanol-acetic acid-water (50:37.5:3.5:2 vol/vol/vol/vol) as previously described (12). After transmethylation,
fatty acid methyl esters from the diphosphatidylglycerol,
glycerophosphocholine, and glycerophosphoethanolamine fractions were
quantitated by capillary gas chromatography (1).
Subcellular fractionation.
Cell extracts were prepared as previously described (16).
Briefly, after fatty acid incubation, cells were grown to near confluence in T-75 or T-175 flasks, trypsinized, pelleted at 200 g, and washed three times with PBS. Subsequently, the cell
pellet was passed through a 27-gauge needle three times with 500 µl
of homogenizing buffer [50 mM Tris · HCl, pH 7.2, 250 mM
sucrose, 2 mM EDTA, 1 mM EGTA, 50 µM sodium fluoride, 100 µM
orthovanadate, 25 µg/ml each of leupeptin, pepstatin, and
aprotinin, and 150 µM 4-(2-aminoethyl)benzenesulfonyl fluoride].
Samples were centrifuged at 100,000 g for 30 min at 4°C,
and the supernatant was taken as the cytosolic extract and frozen in
aliquots at
80°C until use. The pellet was further extracted with
the above buffer supplemented with Triton X-100 at a final
concentration of 1% and then incubated for 30 min on ice. Insoluble
material was removed by a second round of centrifugation at 100,000 g for 30 min at 4°C. The supernatant was saved as the
total membrane extract. In addition, total cell lysates were prepared
with the use of the Triton X-100-containing buffer, incubated on ice
for 30 min, and centrifuged at 14,000 g for 20 min. The
supernatant was designated the total cell extract. Cellular protein was
quantified with the Coomassie Plus assay (Pierce).
Immunoblotting.
Total cellular, membrane, or cytosolic extracts were treated with SDS
sample buffer and subjected to polyacrylamide gel electrophoresis in
4-20% precast mini gels (16). After electrophoresis,
proteins were electroblotted onto a PVDF membrane with the use of a
Hoefer Mighty Small Transphor Unit (Pharmacia, Piscataway, NJ) at 400 mA for 1.5 h. After transfer, the membrane was processed according to the method of Davidson et al. (16), including blocking
of the membrane in 4% nonfat dry milk and 0.1% Tween 20 in PBS at room temperature for 1 h with shaking, followed by incubation with
shaking overnight at 4°C with primary antibody [pan-Ras Ab-3 (Clone
10; Oncogene Science), anti-HMG-CoA reductase (CRL 18811) isolated from
the A9 hybridoma cell line (ATCC, Rockville, MD), or anti-SV40
(Oncogene Science)] diluted in PBS containing 4% milk and 0.1% Tween
20. Membranes were washed with PBS containing 0.1% Tween 20 and
incubated with secondary antibody (peroxidase-conjugated goat
anti-mouse IgG; Kirkegaard & Perry) per manufacturer's instructions. Recombinant H-Ras standard (0.75 ng) from Panvera (Madison, WI) was
used as a marker and resulted in a 21-kDa band. In addition, direct
molecular weight determination of Ras proteins was performed with the
use of positive-ion MALDI-TOF-MS on a PerSeptive Biosystems Voyager
Elite XL system (Framingham, MA) equipped with delayed extraction and a
LSI nitrogen laser operating at 337 nm (4). For HMG-CoA
reductase analysis, mouse brain extract was used as the positive
control. For quantitation, linearity of detection was validated over a
range of sample protein, and blots were scanned and quantitated with a
Fluor-S Max MultiImager System (Bio-Rad, Hercules, CA). Band
intensities were reported as intensity multiplied by band area.
Confocal immunofluorescence.
Cells were plated into four-well pretreated (overnight with 20% FBS in
culture medium) chamber glass slides (catalog no. 136420; Lab-Tek) at a
density of 2,000 or 3,500 cells per well. After 48-72 h, cell
monolayers were rinsed with PBS and fixed with 2% paraformaldehyde-PBS
for 20 min at room temperature (61). Cells were
permeabilized and blocked in PBS containing 0.02% saponin and 5%
normal rabbit serum (SNRS-PBS) for 30 min at room temperature and were
subsequently probed with 20 µg/ml Ras antibody Y13-238 (Oncogene
Science) in SNRS-PBS for 1 h at room temperature. Slides were
washed with PBS containing 0.02% saponin and then incubated with
FITC-conjugated rabbit anti-rat IgG (20 µg/ml; Dako, Carpinteria, CA)
in the dark for 30 min at room temperature, followed by 10 min of
treatment with 100 nM 4',6-diamidine-2-phenylindole dihydrochloride (Roche Molecular Biochemical) to label nuclei. Finally, cells were
washed for 30 min in PBS containing 0.02% saponin and mounted in
ProLong anti-fade mounting medium (Molecular Probes). Slides were
viewed on an Ultima confocal microscope (Meridian Instrument, Okemos, MI).
Farnesyl protein transferase assay.
Farnesyl protein transferase activity was assayed as the ability of
cytosolic protein extracts to catalyze the prenylation of Ras
(16). Cytosolic extracts (40 µg protein) from YAMC-Ras cells were incubated for 1 h with 5 µg of recombinant H-Ras
(Panvera) and 1 µCi of [3H]farnesylpyrophosphate (15.0 Ci/mmol; NEN, Boston, MA) in a buffer containing 50 mM
Tris · HCl, pH 7.5, 10 µM ZnCl2, 20 mM KCl, 5 mM
dithiothreitol (DTT), 0.2% octyl
-D-glucosidase, and
1% DMSO. Samples were precipitated with trichloroacetic acid and
filtered onto glass fiber disks to quantitate transfer of
[3H]farnesylpyrophosphate onto Ras. A range of protein
levels was tested to assure the reaction was proportional to the amount
of protein used. The absence of protein was used as a negative control.
Ras palmitoylation.
After a 54-h pretreatment with 50 µM DHA or LA or with medium alone,
cells were metabolically labeled overnight with 5 mCi/flask of
[35S]methionine (1.25 Ci/mmol; NEN) in RPMI 1640 without
methionine supplemented with 5% FBS, 1% glutamax-1, 1% ITS, 1%
penicillin-streptomycin, and 5,000 U/l IFN-
. After an overnight
incubation, cells were incubated for 4 h with 3.3 mCi/flask of
BSA-complexed [3H]palmitate (49.0 Ci/mmol; NEN) in fatty
acid-free medium to quantify palmitate labeling of Ras
(7). Cell lysates (200-400 µg protein; described in
Subcellular fractionation) were immunoprecipitated by incubating for 16 h at 4°C with 50 µl of pan-Ras antibody
(OP-40, Oncogene Science) and preadsorbed to protein A/G (Oncogene
Research Products, Cambridge, MA). Immunoprecipitates were transferred to spin filters (Cytosignal Research Products, Irvine, CA), rinsed with
PBS, and eluted after a 15-min incubation with SDS-PAGE sample buffer
containing 625 mM Tris · HCl, 2% SDS, 25% glycerol, and 0.001% bromphenol blue (24). Samples were separated by
SDS-PAGE, and gels were exposed to X-ray film for 24 h.
Subsequently, the Ras band (21 kDa) was cut out from the dried gel and
quantitated by dual-channel (3H/35S)
scintillation counting. Counts (dpm) from 35S-labeled Ras
were used as an internal standard in each experiment to normalize the
palmitoylation dpm values (25).
Determination of GTP- and GDP-bound p21ras.
For determination of Ras-GTP and Ras-GDP levels in fatty acid-treated
cells, subconfluent cells were metabolically labeled with 325 µCi/ml
[32P]orthophosphoric acid (285 Ci/mg; NEN) for 4 h
in a low-phosphate medium (23, 44). After
incubation, cells were harvested in a lysis buffer containing 25 mM
Tris · HCl, pH 7.5, 20 mM MgCl2, 144 µM
4-(2-aminoethyl)benzenesulfonyl fluoride, and 1% aprotinin. Samples were centrifuged at 100,000 g for 30 min, and the
supernatant was taken as the cytosolic extract. For preparation of the
membrane fraction, the pellet was resuspended in the above buffer
containing Triton X-100 at a final concentration of 1%, incubated for
30 min on ice, and centrifuged at 100,000 g for 30 min. The
supernatant was saved as the membrane extract. Membrane and cytosolic
extracts were immunoprecipitated with pan-Ras antibody as described in Ras palmitoylation. Ras immunoprecipitates bound to
protein A/G beads were washed with buffer containing 50 mM
Tris · HCl, pH 7.5, 0.2% Triton X-100, 5 mM MgCl2,
500 mM NaCl, and 0.005% SDS (16). Ras immunoprecipitate
was resuspended in 2 mM EDTA, 2 mM DTT, 0.2% SDS, 10 mM GTP, and 10 mM
GDP, as adapted from the procedure of Downward et al.
(18), and incubated for 80 min at 68°C to elute bound
nucleotides. The GTP and GDP from membrane or cytosolic fractions were
resolved by TLC on polyethyleneimine-cellulose plates (Merck),
developed with 2.0 M ammonium formate in 1.0 M HCl as solvent
(40), and detected with a Packard InstantImager scanner.
The percentage of GTP-associated Ras in membrane or cytosolic fractions
was calculated as [dpm GTP/1.5(dpm GTP + dpm GDP)] × 100 (50).
Ras-dependent signal transduction.
To assess the effect of fatty acid treatment on Ras downstream
signaling, cells were incubated with 50 µM LA, DHA, or no fatty acid
under standard culture conditions for 72 h, followed by serum deprivation (0.1% FBS) for 18 h. Select cultures were
subsequently stimulated with 1 µg/ml lipopolysaccharide (Sigma) for
10 min, and total cell lysates were prepared. Activated p42 and p44
extracellular signal-regulated kinase (ERK) levels were determined by
immunoblot analysis with phosphospecific p44/42 ERK antibody, which
detects ERK1 and ERK2 only when catalytically activated by
phosphorylation at Thr-202 and Tyr-204 (Cell Signaling Technology,
Beverly, MA), and p44/42 ERK antibody, which detects total ERK1 and
ERK2 (16). Phosphorylated ERK2 control protein was used as
a standard for ERK blots and yielded a 42-kDa band. Nonphosphorylated
ERK2 was used as a negative control to assess the specificity of the
antibody for the phosphorylated form only. In addition, activated Akt
was determined by immunoblot with a phospho-Akt (Ser-473) antibody, which detects Akt1 when phosphorylated at Ser-473 and Akt2 and Akt3
when phosphorylated at equivalent sites (Cell Signaling Technology). Expression levels of total Akt were also quantitated with the use of
phosphorylation state-independent Akt antibody (Cell Signaling Technology).
Statistical analysis.
Effects of fatty acid treatment on various parameters were analyzed by
one-way ANOVA. Differences (P < 0.05) between means were separated by using the least-squares means test.
 |
RESULTS |
Functional effects of fatty acids on colonocytes in vivo.
We have previously demonstrated that the YAMC cell line is a useful
model system to define the molecular mechanisms by which dietary
constituents modulate colonocyte growth, differentiation, and
apoptosis (5, 19). The YAMC-Ras cell line
overexpresses H-Ras approximately fourfold compared with the parental
cell line (data not shown) and forms visible tumors (15).
Using this model, we first investigated the effects of fatty acids
known to promote (LA, 18:2n-6) and prevent (DHA, 22:6n-3) colon cancer
in experimental animal models (11, 29, 48, 49). After a
72-h incubation, a significant reduction (P < 0.05) in
cell number was observed in 50 µM DHA-treated cultures compared with
LA-treated and control cells at subconfluent densities (Fig.
1). This dose is considered physiologically relevant because it lies well within the range of blood
levels in human subjects supplemented with DHA (14). Fatty
acid treatment had no effect on adherent cell viability (>98%) as
assessed by propidium iodide exclusion. To examine whether growth
arrest by DHA treatment was associated with changes in cell
proliferation and/or apoptosis, we determined BrdU uptake and
caspase 3 activity. DHA treatment significantly reduced DNA synthesis
compared with LA treatment and control (Fig.
2A). In contrast, fatty acid
treatment had no effect on the level of apoptosis in adherent
and floating populations (Fig. 2B). The effects of DHA on
cell proliferation were not the result of alterations in the SV40 gene
product because the expression of SV40 large T antigen was not affected
(data not shown).

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Fig. 1.
Cell number is decreased by docosahexaenoic acid (DHA)
treatment. Young adult mouse colon (YAMC)-Ras cells were treated with
50 µM linoleic acid (LA) or DHA or with medium alone (untreated, UT),
and adherent cells were trypsinized and counted using a hemacytometer
after 72 h. Data are means ± SE from 3 separate experiments
(n = 5). Bars with different symbols represent
treatments that are significantly different (P < 0.05).
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Fig. 2.
Effect of fatty acid treatment on DNA synthesis. Cells
were treated with 50 µM LA or DHA or with medium alone for 72 h.
A: for quantitation of DNA synthesis, 1-2 × 105 cells were pulsed with bromodeoxyuridine (BrdU) 3 h before harvest and processed for BrdU quantitation by ELISA per
manufacturer's instructions. Values are expressed as absorbance at 450 nm. B: the effect of fatty acid treatment on
apoptosis was assessed by quantification of caspase 3 activity
in combined floating and adherent fractions as described in
MATERIALS AND METHODS. Data are mean (±SE) percentages of
untreated control from 3 separate experiments. Bars with different
symbols represent treatments that are significantly different
(P < 0.05).
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Incorporation of fatty acids into membrane phospholipids.
The exogenous fatty acids were differentially incorporated into
membrane phospholipids (Tables
1-3).
DHA was primarily incorporated into the glycerophosphoethanolamine
(EtnGro) and cardiolipin (Ptd2Gro) classes. A significant
(P < 0.05) reduction in arachidonic acid (AA; 20:4n-6)
levels was observed in EtnGro, Ptd2Gro, and
glycerophosphocholine (ChoGro) compared with LA-treated and untreated
cells. Interestingly, compared with untreated cells, DHA-treated cells
exhibited an elevation in the mole-percent level of LA. In contrast, LA
treatment was associated with an elevation in the weight-percent level
of LA in Ptd2Gro and ChoGro and of its metabolic
elongation-desaturation product, AA, in EtnGro compared with
DHA-treated and untreated cells.
n-6 and n-3 PUFA differentially modulate Ras membrane localization.
There were no significant effects of 50 µM fatty acid treatment on
total Ras protein expression (Fig.
3A). Similar to previous reports (13), five- to eightfold more Ras resided at the
membrane compared with the cytosol (Fig. 3B). Interestingly,
the Ras membrane-to-cytosol ratio was 41% higher in LA- vs.
DHA-treated cells (Fig. 3B). When the effects of fatty acid
dose were examined, significant differences (P < 0.05)
between LA and DHA treatments were observed at both the 25 and 50 µM
level (Fig. 4). To corroborate protein
identification by SDS-PAGE, direct molecular weight determination of
the Ras protein was performed with the use of positive-ion MALDI-TOF-MS and was found to be 21,296.8 daltons, consistent with published data
(59). Because Ras must be localized to the inner surface of the plasma membrane to be biologically active (3, 61), we examined its intracellular localization using confocal
immunofluorescence. As expected, Ras was largely targeted to the plasma
membrane (data not shown). Together, these data indicate that fatty
acid treatment can modulate Ras trafficking to the plasma membrane.

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Fig. 3.
Fatty acid treatment alters Ras membrane
localization. A: after a 72-h (50 µM) fatty acid
incubation period, cell lysates (5-10 µg protein,
n = 6-8) were subjected to SDS-PAGE, transferred
to a polyvinylidene difluoride (PVDF) membrane, and probed with a
pan-Ras antibody as described in MATERIALS AND METHODS.
This antibody detects mammalian wild type and activated H-, K-, and
N-Ras proteins. B: cell contents (n = 8-11) were fractionated into membrane (pellet) and cytosolic
(supernatant) compartments by centrifugation at 100,000 g,
and ras intracellular localization was determined by
immunoblotting. Expression was quantified as band intensity (optical
density) multiplied by band area. Data are means ± SE from 5 separate experiments. Bars with different symbols represent treatments
that are significantly different (P < 0.05).
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Fig. 4.
Dose effect of fatty acid treatment on Ras
membrane-to-cytosol ratio. Cells were incubated with fatty acids
(0-100 µM) as described in Fig. 3. , LA
(18:2n-6)-treated cells; , DHA (22:6n-3)-treated cells.
Results are expressed as percentages of untreated control
(n = 3). *Significant difference (P < 0.05) between LA and DHA treatment at 25 and 50 µM levels.
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Fatty acid treatment does not affect Ras posttranslational
processing.
After synthesis, Ras proceeds through a series of posttranslational
steps at the COOH terminus that increase its affinity for the plasma
membrane. The first of these posttranslational modifications is the
addition of a 15-carbon farnesyl lipid at Cys-186. Therefore, we
determined whether the lower Ras membrane-to-cytosol ratio in DHA- vs.
LA-treated cells was due to an effect on farnesylation. As shown in
Fig. 5, no effect on HMG CoA reductase
expression, which produces the farnesylpyrophosphate precursor
mevalonate, was observed after the fatty acid treatment. Similarly,
farnesyltransferase activity was not altered (Fig.
6). Interestingly, although
isoprenylation of Ras is required for cell-transforming activity, it is
posttranslational acylation (palmitoylation of Cys-181 and/or Cys-184)
that directly regulates the amount of protein in the membrane
(25, 26). However, fatty acid treatment had no effect on
the relative ratio of [3H]palmitate- to
[35S]methionine-labeled Ras, an index of palmitoyl
transferase activity in vivo (Fig. 7).
Although farnesylation activity and HMG-CoA reductase expression are
indirect markers of farnesylation status, without isoprenoid attachment
to Cys-186 the H-Ras protein remains soluble, and the adjacent Cys-181
and Cys-184 do not become palmitoylated (3, 7, 37).
Therefore, any perturbation in farnesylation would negatively impact
palmitoylation, the terminal processing step. Because fatty acid
treatment had no effect on Ras palmitoylation status, we conclude that
posttranslational processing was not significantly influenced by fatty
acid treatment.

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Fig. 5.
Colonocyte HMG CoA reductase expression. Cells were
treated with 50 µM LA or DHA or with medium alone for 72 h.
Membrane (100,000 g pellet) extracts (20 µg of protein)
were immunoblotted as described in MATERIALS AND METHODS.
Expression was quantified as band intensity (optical density)
multiplied by band area. Data are mean (±SE) percentages of untreated
control from 3 separate experiments (n = 5-11).
There were no significant differences among treatment groups
(P > 0.05).
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Fig. 6.
Farnesyl protein transferase activity 72 h after
fatty acid treatment. Cells were treated with 50 µM LA or DHA or with
medium alone for 72 h. Cytosolic (100,000 g
supernatant) extracts (40 µg) were incubated for 1 h at 37°C
with [3H]farnesylpyrophosphate (0.5 µCi/µl) and 5 µg of recombinant H-Ras. Enzyme activity was assayed as counts (dpm)
of [3H]farnesylpyrophosphate transferred per 40 µg of
protein per hour. Data are means ± SE from 3 separate experiments
(n = 5-6). No significant differences among
treatment groups were noted (P > 0.05).
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Fig. 7.
Lack of an effect of fatty acid treatment on the ratio of
[3H]palmitate- to [35S]methionine-labeled
Ras, an index of palmitoyl transferase activity. Cells were treated for
72 h with 50 µM DHA or LA or with medium alone. During the final
24 h of fatty acid treatment, cells were metabolically labeled
overnight (16 h) with [35S]methionine (250 µCi/ml) in a
low-methionine medium, followed by a 4-h incubation with
[3H]palmitic acid (165 µCi/ml). After fatty acid
incubation, cells were harvested and Ras immunoprecipitated with
pan-Ras antibody. Immunoprecipitates were subjected to SDS-PAGE, and
the 21-kDa band was excised and dual counted for 3H and
35S as described in MATERIALS AND METHODS.
Counts (dpm) from 35S-labeled Ras were used as an internal
standard in each experiment to normalize the palmitoylation dpm values.
Data are means ± SE from 3 separate experiments
(n = 7-8). There were no significant differences
among treatment groups (P > 0.05).
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DHA suppresses Ras activation.
Because localization of Ras to the plasma membrane and its subsequent
binding to GTP are required steps for activation and downstream
signaling, the percentage of Ras bound to GTP in membrane and cytosol
fractions was assessed. A significant reduction (P < 0.05) was observed in the percentage of GTP bound Ras in the membrane
fraction of DHA-treated cells compared with LA-treated and untreated
cells (Fig. 8A). In contrast,
no significant differences were observed in the percentage of GTP bound
Ras in the cytosolic fraction (Fig. 8B). As expected, the
majority (89%) of GTP bound Ras was localized at the membrane (Fig.
8C).

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Fig. 8.
DHA incubation suppresses membrane Ras GTP-binding. Fatty
acid-treated cells (50 µM for 72 h) were incubated with 325 µCi/ml [32P]orthophosphoric acid in phosphate-free
medium for 4 h. After incubation, cells were harvested and
fractionated into 100,000 g membrane and cytosolic
compartments as described in MATERIALS AND METHODS. Ras
immunoprecipitates were subjected to thin-layer chromatography to
separate nucleotide classes. Radiolabel was quantitated with an
InstantImager scanner to determine the percent GTP- vs. percent
GDP-bound Ras. A: membrane Ras GTP levels (n = 9). B: cytosolic Ras GTP levels (n = 9).
C: total GTP-bound Ras levels, membrane vs. cytosol
(n = 27). Data are means ± SE from 3 separate
experiments. Bars with different symbols represent treatments that are
significantly different (P < 0.05).
|
|
Ras activation can lead to the sequential stimulation of ERK;
therefore, expression of activated ERK was quantified by using a
phosphorylated ERK-specific antibody. Because it is difficult to
quantify the activation of many signaling molecules under
"nonstimulated/basal" growth conditions, we attempted to highlight
the differential induction of ERKs using a potent inflammatory stimuli,
i.e., lipopolysaccharide (36, 42). The level of activated
p42 and p44 ERK was 63% lower in DHA-treated cells compared with
LA-treated cells (Fig. 9). In addition,
there was no difference in total ERK expression (phosphorylated plus
nonphosphorylated) in any of the treatments (data not shown). We next
examined the ability of DHA to modulate the activation of a second Ras
effector, i.e., the phosphatidylinositol (PI) 3-kinase/Akt signaling
pathway. Data from these experiments indicated no effect of fatty acid
treatment on either the level of activated Akt or total Akt, i.e.,
phosphorylated and nonphosphorylated (data not shown). These data
suggest that PUFA modulate the ERK signaling pathway.

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|
Fig. 9.
DHA incubation suppresses lipopolysaccharide-induced
extracellular signal-regulated kinase (ERK)1/ERK2 activation. After
fatty acid incubation (50 µM) for 72 h, cultures were serum
deprived for 18 h and stimulated with lipopolysaccharide (1 µg/ml) for 10 min. Cell lysates were prepared, and activated p42/44
ERK levels were determined by immunoblotting with phosphospecific ERK
antibody as described in MATERIALS AND METHODS. Data are
means ± SE from 2 separate experiments (n = 3-5). Bars with different symbols represent treatments that are
significantly different (P < 0.05).
|
|
 |
DISCUSSION |
At present, the molecular basis for the protective effect of fish
oil on the incidence of colon cancer is a complete black box. In a
previous study, we demonstrated that dietary fish oil modulates Ras
intracellular localization by reducing its levels at the plasma
membrane in colonocytes isolated from rats injected with carcinogen
(16). This was associated with a suppression of colonic
tumor development (10, 11). In the present study, we
conducted a series of complementary experiments to investigate the
mechanism(s) by which fish oil decreases Ras membrane localization. Specifically, we examined the biological properties of DHA
(antitumorigenic), the major n-3 PUFA found in fish oil, compared with
those of LA (protumorigenic), the major n-6 PUFA found in corn oil, in
a mouse malignant transformed colonic cell line that overexpresses
oncogenic H-Ras (YAMC-Ras). Because posttranslational processing is
essential for Ras membrane anchoring, we examined whether purified
fatty acids found in fish oil and corn oil affect protein farnesylation and/or palmitoylation. In addition, for the purpose of contrasting PUFA
treatments, a third "control" group was supplemented with fatty
acid-free BSA only. It is important to note, as a caveat to this
approach, that uncomplexed BSA can serve as a fatty acid sink and
therefore also can exert a biological effect distinct from fatty acid
supplementation (54). Our data demonstrate that although
the Ras membrane-to-cytosol ratio, an indication of activation state,
was lower in DHA-treated cells (4.9) than in LA-treated cells (8.3),
Ras farnesylation and palmitoylation were not affected. These data are
consistent with our rat colon data (16) and indicate that
long-chain PUFA do not modulate Ras posttranslational processing. Although the mechanism(s) responsible for these observations has not
been elucidated, it is likely that distinct classes (n-3 vs. n-6) of
PUFA can alter membrane structure, thereby influencing the interaction
of Ras proteins with specific domains within the plasma membrane
(58). Interestingly, compared with the untreated control,
it appears as though LA treatment actually increases Ras membrane
association. However, this type of contrast is difficult to
interpret because, in vivo, there is no comparable control diet.
Ras activation is dependent on its binding to GTP (30).
Ras is predominantly in the GDP-bound state, and upon interaction with
the Grb2-SOS complex at the plasma membrane, GTP is exchanged for GDP
(21, 50). Subsequently, it is cycled back to the GDP state
by GTPase-activating protein (GAP) activity (56). Select fatty acids may affect the GTP/GDP binary switch by modulating GAP
activity and function, which regulates hydrolysis of GTP to GDP and,
hence, the Ras activation state (6, 21, 22).
Interestingly, investigators have shown that AA is capable of
inhibiting GAP activity (53, 62). Therefore, we further
investigated the effect of DHA and LA (an AA precursor) on the fatty
acid composition of membrane phospholipid classes. It is evident that
DHA treatment enriches membrane phospholipid levels of DHA and
eicosapentaenoic acid (EPA) and reduces AA, indicating that exogenous
PUFA are incorporated into membrane lipids. This is consistent with
previously published data where fish oil was fed to rats and was shown
to decrease the AA content of colonic epithelial cell total
phospholipids (35). The antagonism of AA by DHA is
noteworthy in view of the documented inhibitory effect of AA on Ras
GAPs (28, 53, 57). Because GAPs deactivate Ras, the
anticipated outcome of a suppression of membrane AA content would be a
decrease in the levels of GTP-bound (active) Ras in colonocytes. This
is precisely what was observed in cells treated with DHA.
It is well established that in the active, GTP-bound state, Ras can
activate the mitogen-activated protein kinase/ERK kinase pathway
(9). Activated ERK kinases can translocate to the
nucleus and regulate the activity of transcription factors such as
Elk-1 (40). Membrane GTP-bound Ras additionally interacts
with and activates the heterodimer enzyme PI 3-kinase
(41). Increased PI 3-kinase activity results in the
activation of Akt, a serine/threonine kinase (32, 41) that
promotes cell survival (33). To further examine the
influence of DHA on Ras activation and effector protein function,
p42/44ERK activation was determined by phosphospecific
immunoblotting after lipopolysaccharide stimulation. ERK activation was
reduced by 63% in DHA- vs. LA-treated cells. Interestingly, DHA
treatment did not block Akt activation, which is consistent with a lack of effect on the level of apoptosis. Collectively, these data indicate that DHA disrupts Ras-dependent signal transduction by suppressing plasma membrane localization, GTP binding, and
p42/44ERK activation. This may partly explain why dietary
fish oil protects against colon cancer development.
In conclusion, we have demonstrated that DHA (isolated from fish oil),
when compared with LA (isolated from corn oil), lowers the activation
of the Ras oncogene by 1) reducing Ras localization to the
plasma membrane without affecting posttranslational processing, 2) suppressing levels of GTP-bound (activated) Ras at the
plasma membrane, and 3) partially blocking downstream signal
transduction. These observations are relevant because they corroborate
previously published in vivo findings (16). In these
studies, we demonstrated that dietary fish oil compared with corn oil
downmodulates Ras intracellular localization by reducing its levels at
the plasma membrane in colonocytes isolated from rats injected with
azoxymethane (a colon carcinogen). This was associated with a
suppression of colonic tumor development (10) and altered
crypt cytokinetics (29). Because the in vivo vs. in vitro
effects of DHA and LA on Ras intracellular localization are comparable,
we propose that YAMC-Ras cells are a suitable model system to probe
unresolved mechanistic questions. Further experiments with the use of
quantitative immunofluorescence are required to determine the effect of
PUFA classes on the precise intracellular localization of specific isoforms of Ras. In addition, whether other prenylated G proteins such
as Rho GTPases are modulated by select PUFA merits further investigation.
 |
ACKNOWLEDGEMENTS |
We thank Nancy Turner for statistical analyses, Lloyd Sumner for
mass spectrometry, Rola Barhoumi and Robert Burghardt for confocal
microscopy, and Jianhu Zhang for technical assistance.
 |
FOOTNOTES |
This work was supported in part by National Cancer Institute Grants
CA-59034 and CA-61750, National Institute of Environmental Health
Sciences Grant P30-ES-09106, and American Institute for Cancer Research
Grant 00A055.
Address for reprint requests and other correspondence:
R. S. Chapkin, 442 Kleberg Biotechnology Center, 2471 TAMU, Texas
A&M Univ., College Station, TX 77843-2471 (E-mail:
r-chapkin{at}tamu.edu).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 21 July 2000; accepted in final form 7 November 2000.
 |
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