Differential effects of eicosapentaenoic acid and docosahexaenoic acidin promoting the differentiation of 3T3-L1 preadipocytes

Ganesan Murali a,b,c, Cyrus V. Desouza c,a, Michelle E. Clevenger a,c, Ramesh Ramalingam a,c,Viswanathan Saraswathi a,b,c,n

a Departments of Internal Medicine, Division of Diabetes, Endocrinology, and Metabolism, Omaha, NE, United Statesb Cellular and Integrative Physiology, University of Nebraska Medical Center, Omaha, NE, United Statesc Research Services, VA Nebraska Western Iowa Health Care System, Omaha, NE, United States

a r t i c l e i n f o

Article history:Received 4 July 2013Received in revised form1 October 2013Accepted 30 October 2013

Keywords:n-3 fatty acidsAdipogenesisEPADHADocosapentaenoic acid

a b s t r a c t

The objective of this study was to determine the effects of enrichment with n-3 fatty acids,eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA), on the differentiation of 3T3-L1preadipocytes. Enrichment with DHA but not EPA significantly increased the differentiation markerscompared to control differentiated cells. DHA compared to EPA treatment led to a greater increase inadiponectin secretion and, conditioned media collected from DHA treated cells inhibited monocytemigration. Moreover, DHA treatment resulted in inhibition of pro-inflammatory signaling pathways. DHAtreated cells predominantly accumulated DHA in phospholipids whereas EPA treatment led toaccumulation of both EPA and its elongation product docosapentaenoic acid (DPA), an n-3 fatty acid.Of note, adding DPA to DHA inhibited DHA-induced differentiation. The differential effects of EPA andDHA on preadipocyte differentiation may be due, in part, to differences in their intracellular modificationwhich could impact the type of n-3 fatty acids incorporated into the cells.

& 2013 Elsevier Ltd. All rights reserved.

1. Introduction

Obesity, a feature of metabolic syndrome, predisposes indivi-duals to type 2 diabetes and cardiovascular disease. Adipose tissue(AT) dysfunction, characterized by adipocyte hypertrophy, insulinresistance, and increased lipolysis, is considered to mediate thepathological consequences of obesity [1,2]. In particular, impairedinsulin sensitivity in AT leads to increased basal lipolysis therebyleading to excessive release of free fatty acids (FFAs) (reviewed in[3]). These FFAs play an important role in the development ofobesity-associated metabolic disorders including insulin resis-tance, dyslipidemia and hepatic steatosis [2–4]. Thus, properstorage of lipids in the AT is essential to prevent lipotoxicitycaused by FFAs.

It is well established that a dysfunctional AT not only exhibitsexcessive lipolysis but also secretes pro-inflammatory cytokines andchemokines. It should also be pointed out that AT in obesity exhibitsa unique phenotype characterized by the accumulation of inflam-matory cells, predominantly macrophages. The AT macrophagesplay an important role in the secretion of pro-inflammatory

mediators from obese AT [5,6]. In fact, the accumulation of macro-phages in the AT is preceded by the development of insulinresistance (IR) in mouse models [5].

Dietary fish oil containing n-3 fatty acids such as eicosapentae-noic acid (EPA) and docosahexaenoic acid (DHA) is known toregulate both the storage and secretory functions of AT (reviewedin [7]). We previously showed that fish oil increased visceral ATmass which was associated with several beneficial metaboliceffects in low density lipoprotein receptor deficient mice [8].Moreover, Ide has shown that fish oil increased visceral AT masswith a concurrent decrease in hepatic steatosis and dyslipidemiain ICR mice [9]. In other studies, fish oil has been shown todecrease or not to alter adiposity. Overall, the effect of fish oil onadiposity appears to depend on the mouse strain and otherexperimental conditions. Regardless of its effects on adiposity, fishoil has been shown to improve the secretory functions of AT and toreduce AT-specific inflammation [7]. Taken together, these variousstudies show that the n-3 fatty acids found in fish oil have thepropensity to alter the storage and/or the secretory functionsof AT.

A few studies have addressed the direct effects of n-3 fattyacids in modulating the differentiation of preadipocytes intoadipocytes in vitro but the results are inconclusive. For example,EPA has been shown to inhibit, promote, or not to alter thedifferentiation of preadipocytes [10–12]. On the other hand, DHA

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Prostaglandins, Leukotrienes and EssentialFatty Acids

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n Corresponding author at: Research Services, VA, Nebraska Western Iowa HealthCare System, Omaha, NE. Tel.: þ1 402 995 3033; fax: þ1 402 449 0604.

E-mail address: s.viswanathan@unmc.edu (V. Saraswathi).

Prostaglandins, Leukotrienes and Essential Fatty Acids 90 (2014) 13–21

can promote or inhibit the differentiation of preadipocytes intoadipocytes [11,13]. Thus, the effects of EPA versus DHA enrichmentin modulating preadipocyte differentiation are still unclear.

It is becoming widely recognized that although both EPA andDHA are n-3 fatty acids, they have differential effects in modulat-ing the cellular functions. For example, DHA but not EPA reducedTNFα-induced inflammatory response in human umbilical veinendothelial cells [14]. Also, DHA is more effective than EPA inreducing TNFα-induced cell death [15]. Conversely, EPA is morepotent than DHA in attenuating LPS-induced inflammatoryresponse in human alveolar macrophages [16]. Therefore, thequestion as to whether these two fatty acids are equivalent ordifferent still remains unresolved. Regardless of their specificeffects in various cell types, a plethora of literature suggests thebeneficial effects of both of these fatty acids in amelioratingmetabolic and cardiovascular disorders (reviewed in [17,18]).Understanding the specific mechanisms by which these fatty acidsmodulate adipocyte functions would be helpful to maximize theirbenefits in ameliorating diseases associated with obesity. There-fore, the objective of this study was to determine the effects of EPAversus DHA on preadipocyte differentiation and to elucidate thepotential mechanisms by which these two fatty acids modulatethe differentiation process.

We used 3T3-L1 preadipocytes and pretreated them with n-3fatty acids individually or in combination and compared the effectsof these fatty acids with oleic acid (OA), a mono-unsaturated fattyacid. We have demonstrated that DHA was more potent ininducing the differentiation of 3T3-L1 preadipocytes comparedto EPA. DHA-induced preadipocyte differentiation was associatedwith improved adipocyte secretory functions as evident fromincreased adiponectin release and reduced monocyte migrationtowards adipocyte conditioned media. Finally, we show that EPAand DHA treatment led to differential incorporation of n-3 fattyacids in phospholipids which could partly account for the differ-ential effects of these fatty acids on preadipocyte differentiation.

2. Materials and methods

2.1. Differentiation of 3T3-L1 preadipocytes

3T3-L1 preadipocytes were originally purchased from Amer-ican Type Culture Collection and were kindly provided byDr. Robert Bennett (VA Nebraska Western Iowa Health CareSystem/University of Nebraska Medical Center, Omaha, NE).On Day 1, the cells were cultured in Dulbecco's Modification ofEagle's Medium (DMEM) (Mediatech, Manassas, VA) supplemen-ted with 10% bovine calf serum (BCS) (Life Technologies, GrandIsland, NY). We used the same batch of BCS in all our experiments.The fatty acid treatment was carried out using media containingserum as the source of albumin as we described previously [19].Briefly, the fatty acids were first dissolved in ethanol and wereadded to DMEM containing 10% serum before treating the 3T3-L1cells. The cells were pretreated for 24 h (Day 2) with differentfatty acids at 50 mM concentration (Nu Chek Prep, Elysian, MN). Forcombined treatment with EPA and DHA at 50 mM concentration,these fatty acids were used at a ratio of 1.5:1 (EPA 30 mM and DHA20 mM) to reflect the fatty acid composition of the commerciallyavailable fish oil supplements often used in clinical settings.Differentiation of 3T3-L1 preadipocytes into adipocytes wasinitiated by the addition of 250 nM dexamethasone, 0.5 mMisobutyl methylxanthine, and 167 nM insulin (Day 3). Cells wereallowed to differentiate in the presence or absence of fatty acidsfor 48 h. The cells were then maintained in medium containing167 nM insulin in the presence or absence of fatty acids (Day 5) fortwo more days after which the cells and media were collected for

various analyses (Day 7). Basically, we performed all our experi-ments 4 days post-treatment with differentiation media (2 dayswith differentiation media followed by 2 days with insulin-containing media), in the presence or absence of fatty acids.

2.2. Bodipy and oil red O staining

Lipid accumulation in cells at the end of differentiation (Day 7)was analyzed by Bodipy staining. The cells were washed threetimes with PBS and incubated with 10 mM Bodipy (Invitrogen) inPBS for 1 h at room temperature. Cells were washed and, cover-slipped with prolong gold anti-fade reagent containing DAPI(Invitrogen). We also quantified the amount of neutral lipids afterstaining the cells with oil red O as reported previously [20]. Thecolor intensity was measured at 490 nm in a spectrophotometer.

2.3. Realtime PCR analysis for gene expression

The 3T3-L1 preadipocytes were pretreated with fatty acids anddifferentiated as described above. At the end of the differentiationprocess (Day 7), the cells were harvested in trizol and the totalRNA was isolated and real-time PCR was carried out as wedescribed earlier [21]. We used the Taqman realtime PCR primerprobes from Applied Biosystems for the gene expression analyses.We used 18s rRNA as an internal control.

2.4. Western blot

At the end of differentiation (Day 7), the cell lysates werecollected and western blot analysis was carried out for themembers of the mitogen-activated protein kinase (MAPK) signal-ing pathway including extracellular signal related kinase (ERK1/2),p38, and c-jun NH2-terminal kinase/stress-activated proteinkinase (JNK/SAPK). Glyceraldehyde 3-phosphate dehydrogenase(GAPDH) was used as an internal control (Cell Signaling Tech).Additionally, western blot analysis was performed in conditionedmedia collected after the differentiation process (Day 8) foradiponectin, an adipocyte-derived secretory factor. Briefly, wechanged the insulin containing medium at the end of the differ-entiation process (Day 7) and replaced it with regular growthmedium (DMEMwith 10% BCS) with or without various fatty acids.We then collected the conditioned media after 24 h (Day 8).

2.5. Boyden chamber assay for monocyte migration

Human THP-1 monocytes were originally purchased from ATCCand maintained in culture in RPMI 1640 containing 10% fetalbovine serum. Migration studies were performed using an MBSeries 96-well Boyden chamber (Neuroprobe). The top and bottomchambers were separated by a polycarbonate filter with 8-μmpores. The bottom wells were loaded with 390 μl of conditionedmedia collected as described above. The top wells were loadedwith 100 μl of THP-1 cells at a concentration of 3�106 cells/mlfollowed by incubation at 37 1C for 2 h. THP-1 monocytes thatmigrated to the bottom chamber were stained with 0.4% trypanblue and counted using the Countless automated cell counter(Invitrogen). We counted both live and dead cells in the bottomchamber at the end of the migration assay.

2.6. Gas chromatographic analysis for fatty acid profilein phospholipids

We analyzed the content of n-3 fatty acids in phospholipidsby the gas chromatographic method as we described previously[22]. Briefly, at the end of differentiation (Day 7), cells were rinsedtwice with PBS followed by another wash using 0.1% BSA in PBS.

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The cells were scraped in PBS followed by centrifugation to collectthe cell pellet. Lipids were extracted using the method ofFolch et al. [23]. Individual lipid classes were separated by thinlayer chromatography using Silica Gel 60A plates developed inpetroleum ether–ethyl ether–acetic acid (80:20:1) and visualizedby rhodamine 6G. Phospholipids were scraped from the platesand methylated using BF3/methanol as described by Morrisonand Smith [24]. The methylated fatty acids were extractedand analyzed by gas chromatography using an HP 7890A gaschromatograph equipped with a SP-2380 capillary column(0.25 mm�30 m, 0.25 μm film) and a flame ionization detector.Fatty acid methyl esters were identified by comparing the reten-tion times with those of known standards. Odd chain fatty acidswere used as internal standards to quantify the amount of lipid inthe sample.

2.6.1. Statistical analysisValues are presented as the mean7SEM. Data were analyzed

with Prism Graphpad software. One way analysis of variancefollowed by Tukey's post-hoc test was performed to compare theresponses among different groups. A statistical probability ofPo0.05 was considered significant.

3. Results

3.1. DHA was more potent than EPA in inducing the differentiationprocess in 3T3-L1 preadipocytes

To determine the effects of n-3 fatty acids in modulatingpreadipocyte differentiation, we pretreated cells with these fattyacids followed by adding the differentiation media in the presenceor absence of fatty acids. After two days treatment with differ-entiation medium followed by two days treatment in insulincontaining medium, we stained the cells with Bodipy, a fluores-cent dye which stains lipid droplets. As shown in Fig. 1, the cellstreated with DHA showed a greater increase in differentiationcompared to other fatty acids (Fig. 1A–F). We next quantified theamount of lipids present in each group by staining the lipiddroplets with oil red O. We noted that oil red O staining issignificantly increased in all groups compared to undifferentiatedcells. The cells pretreated with EPA, DHA, or EþD showed asignificant increase in oil red O staining compared to differentiatedcontrol. However, cells pretreated with DHA showed a muchgreater increase in oil red O staining. Interestingly, we noted thatoil red O staining was significantly higher in DHA (Po0.001)compared to EPA treated cells. Furthermore, adding EPA to DHA(EþD) significantly reduced oil red O staining compared to DHAalone treated cells (Fig. 1G).

3.2. DHA significantly increased the mRNA expressionof differentiation markers

We performed realtime PCR analysis to analyze the expressionof genes involved in cell differentiation (Fig. 2A–C). We noted thatthe cells treated with DHA showed a marked increase in theexpression of CCAAT/enhancer-binding protein alpha (C/EBPα),peroxisome proliferator activated receptor gamma (PPARγ), andadipocyte protein 2 (aP2). C/EBPα and PPARγ mRNA expressionwas significantly increased in all groups compared to undiffer-entiated control. However, a significant increase was noted only incells receiving EPA, DHA or EþD compared to differentiatedcontrol. Notably, C/EBPα and PPARγ mRNA levels were signifi-cantly higher in DHA even when compared to EPA and EþDtreated cells. Regarding aP2 mRNA expression, a downstreamtarget of PPARγ and a marker of mature adipocytes, a significant

increase was noted in all groups compared to undifferentiatedcontrol. However, only cells treated with DHA and EþD but notEPA showed a significant increase compared to differentiatedcontrol. Taken together, DHA was more potent compared to EPAin inducing preadipocyte differentiation and adding EPA canactually blunt markers of differentiation induced by DHA.

3.3. DHA is more potent than EPA in increasing adiponectin secretion

We next analyzed the conditioned media for adiponectinprotein levels by western blot analysis, and we noted thatadiponectin is increased significantly upon differentiation in allthe groups. We also noted that DHA was the most potent of all inincreasing adiponectin release (Fig. 3A and B). In particular,adiponectin level was significantly higher in conditioned mediaderived from DHA versus EPA treated cells (Po0.01).

3.4. Conditioned media collected from 3T3-L1 preadipocytesdifferentiated in the presence of DHA reduces monocyte migration

Macrophage accumulation in AT is often associated with ATinflammation and other pathophysiological consequences of obe-sity. We performed the cell migration experiment using condi-tioned media at the bottom chamber and THP-1 monocytes at thetop chamber. We noted that media collected from OA and EPAtreated cells showed a trend towards a decrease in monocytemigration compared to both controls. Interestingly, only the mediacollected from DHA treated cells showed a significant decrease inmonocyte migration compared to undifferentiated and differen-tiated controls (Fig. 3C). Moreover, the cell migration was sig-nificantly decreased by media from DHA compared to EPA treatedcells. These data indicate that in addition to promoting differentia-tion, DHA treatment leads to improved secretory functions ofadipocytes thereby leading to reduced monocyte migration.

3.5. DHA inhibits signaling pathways altering inflammationand/or adipogenesis

Since the n-3 fatty acids are potent anti-inflammatory agents, wenext wanted to study their effects in altering the pro-inflammatorysignaling pathways, in particular, MAPKs (Fig. 4A–D). Of note, MAPKsare also involved in regulating adipogenesis. As shown in Fig. 4B, thecells differentiated in the presence of OA and EPA showed anincrease in ERK1/2 phosphorylation compared to undifferentiatedcontrol cells. Interestingly, the cells differentiated in the presence ofDHA exhibited a significant decrease in ERK1/2 activation comparedto differentiated control cells and cells differentiated in the presenceof EPA. We next analyzed p38 MAPK, another member of MAPKs.We noted a significant increase in p38 MAPK in EPA treated cellscompared to undifferentiated control. However, p38 MAPK activa-tion was significantly decreased in DHA treated cells compared todifferentiated control and EPA groups (Fig. 4C). The phosphorylationof JNK/SAPK was not altered significantly in any of the groups(Fig. 4D). Taken together, these data suggest that EPA and DHA exertdifferential effects in modulating the activation of MAPKs and thatDHA induced preadipocyte differentiation is associated with reducedactivation of pro-inflammatory signaling pathways, in particular,ERK1/2 and p38 MAPKs.

3.6. Enrichment with EPA versus DHA differentially modulatesthe pattern of n-3 fatty acids in phospholipids

Since n-3 fatty acids can modulate membrane fatty acidcomposition [25], and because phospholipids are an integral partof cell membranes, we studied the accumulation of n-3 fatty acidsin phospholipids. As expected, our data showed that EPA and DHA

G. Murali et al. / Prostaglandins, Leukotrienes and Essential Fatty Acids 90 (2014) 13–21 15

treatment led to a profound increase in the accumulation of EPAand DHA, respectively, in phospholipids (Fig. 5). Surprisingly, wealso noted that EPA treatment not only increased EPA but also DPA,

an elongation product of EPA. Notably, DHA treatment led to apronounced increase in DHA accumulation. Moreover, the EPAcontent is significantly increased with a concurrent decrease in the

Fig. 1. Effects of n-3 fatty acids on the differentiation of 3T3-L1 preadipocytes. 3T3-L1 preadipocytes were pretreated and differentiated in the presence or absence of variousfatty acids. Bodipy staining was performed to detect lipid droplets and fluorescent images were obtained at 20� magnification (A–F). In separate experiments, after thedifferentiation process, the cells were stained with oil red O. The lipid droplets were dissolved in 60% isopropanol and the color intensity was measured at 490 nm in aspectrophotometer. Values are expressed as mean7SEM of 9 samples in each group (G). UD, undifferentiated control; Diff, differentiated control; OA, oleic acid; EPA,eicosapentaenoic acid; DHA, docosahexaenoic acid; and EþD, EPAþDHA.

Fig. 2. Effects of n-3 fatty acids on the mRNA expression of differentiation markers. RNA samples were analyzed by real time PCR for the expression of genes involved in thedifferentiation of 3T3-L1 preadipocytes (A–C). Values are expressed as mean7SEM of 18 samples in each group. C/EBPα, CCAAT/enhancer-binding protein alpha; PPARγ,peroxisome proliferator activated receptor gamma; aP2, adipocyte protein 2.

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DPA content in DHA treated cells compared to both undifferen-tiated and differentiated controls. This suggests that the intracel-lular processing of EPA and DHA leads to differential accumulationof n-3 fatty acids in phospholipids.

3.7. The pro-adipogenic effects of DHA is impaired in the presenceof DPA

The GC analysis revealed that DPA, an elongation product ofEPA, represents about 33% of total very long chain n-3 fatty acidscontent in EPA treated cells. Therefore, it is reasonable to speculate

that the smaller magnitude of differentiation seen in EPA com-pared to DHA treated cells could be due to its conversion to DPA.To check this possibility, we differentiated cells with DHA in thepresence or absence of DPA. As shown in Fig. 6A, lipid accumula-tion was significantly increased in all the groups compared toundifferentiated control. Only DHA showed a significant increasein lipid accumulation compared to differentiated control. Notably,the extent of lipid accumulation was not altered in DPA treatedcells compared to differentiated control. Further, we noted thatadding DPA to DHA significantly reduced DHA-induced lipidaccumulation compared to DHA alone treated cells. Moreover,markers of differentiation were reduced upon adding DPA to DHA(Fig. 6B–D).

4. Discussion and conclusions

In the current investigation, we have compared the effects ofEPA versus DHA in modulating the differentiation of preadipocytesto adipocytes. We have demonstrated that DHA is more potentthan EPA in inducing the differentiation process. We provide novelevidence that while DHA treatment led to a predominant increasein DHA accumulation in phospholipids, EPA enrichment resulted inaccumulation of both EPA and DPA. Further, we have shown thatadding DPA to DHA reduced its adipogenic effect. Taken together,our data suggest that the differential effects of EPA and DHA onpreadipocyte differentiation may be partly mediated via differen-tial accumulation of n-3 fatty acids in membrane phospholipids.

The effect of n-3 fatty acids in modulating the differentiationprocess is still unclear. Hanada et al. showed that EPA was effectivein promoting the differentiation process [11]. Tanabe et al. showedthat adding EPA during the differentiation of 3T3-L1 cells did notalter the lipid content but significantly reduced PPARγ2, a markerof differentiation [12]. In another study, Manickam et al. showedthat EPA at 100 mM leads to reduced lipid accumulation andreduced mRNA expression of PPARγ [10]. Our data show thatEPA by itself induced preadipocyte differentiation to some extent.However, adding EPA to DHA resulted in diminished adipogenesis.

Regarding the role of DHA in modulating adipogenesis, Tanabeet al. showed that adding this fatty acid during preadipocytedifferentiation did not alter lipid accumulation but reduced themRNA expression of PPARγ2 [12]. Kim et al. showed that DHA hasanti-adipogenic effect in 3T3-L1 preadipocytes [13]. In contrast,our study showed that DHA is very potent in inducing thedifferentiation process. One major difference among these studiesis that Tanabe et al. and Kim et al. differentiated cells in thepresence of DHA; whereas we first enriched 3T3-L1 cells with fattyacids for 24 h followed by differentiation in the presence of fattyacids. This might have resulted in differential activation of signal-ing pathways involved in preadipocyte differentiation. In particu-lar, MAPK/ERK activation has been shown to inhibit preadipocytedifferentiation [26,27]; whereas inhibition of this pathway canpromote differentiation [28,29]. In line with these findings, wenoticed a significant decrease in ERK1/2 phosphorylation in cellsdifferentiated in the presence of DHA and these data suggest thatinhibition of ERK1/2 activation is one mechanism by which DHApretreatment promotes differentiation in our study. It should alsobe pointed out that the role of ERK1/2 in modulating adipogenesisis debatable. Some studies have shown that an increase in ERK1/2phosphorylation can promote preadipocyte differentiation [30,31].Kim et al. have reported that the timing of ERK1/2 activation iscritical in modulating the differentiation process. For example,they have shown that inhibition of ERK1/2 on Day 2 after initiatingthe differentiation process promotes differentiation [26]. Also, itwas hypothesized that initially, ERK has to be turn on for aproliferative step, while latter on, it has to be shut-off to avoid

Fig. 3. Effects of n-3 fatty acids on the secretory functions of adipocytes. 3T3-L1cells were differentiated in the presence or absence of various fatty acids. After thedifferentiation process (Day 7), the cells were washed and incubated with DMEMwith 10% BCS in the presence or absence of respective fatty acids for an additional24 h (Day 8). The media samples were collected on Day 8 and analyzed for theprotein levels of adiponectin by Western blot analysis (A, B). A representative bandfrom each group is shown. Values are expressed as mean7SEM of 6 samples ineach group. Cell migration was analyzed by in vitro Boyden chamber migrationassay (C). The conditioned media collected as described above were added to thelower compartment. THP-1 monocytes were added into upper compartment andthe chamber was incubated at 37 1C for 2 h and the number of transmigrated cellsin the bottom chamber was scored. Values are mean7SEM of 18 samples ineach group.

G. Murali et al. / Prostaglandins, Leukotrienes and Essential Fatty Acids 90 (2014) 13–21 17

PPARγ phosphorylation and to promote adipogenesis (reviewedin [32]). As mentioned, we first pretreated the cells with variousfatty acids followed by differentiation. Thus, when we initiated the

differentiation process, the cells already reached confluence andinhibition of ERK1/2 by DHA at this time might have resulted in anenhanced differentiation. Our findings are further supported bythe observation that DHA can induce the differentiation of AML1cells which are preadipocytes of human origin [11]. In this study,Hanada et al. showed that DHA promotes the differentiationprocess however at a higher concentration (250 mM).

Obesity-associated AT inflammation is characterized byincreased macrophage accumulation in AT. We and others haveshown that fish oil can reduce AT macrophage accumulation[8,33]. Our current data are in agreement with the widely acceptednotion that the n-3 fatty acids are anti-inflammatory. First, weshow that the conditioned medium collected from DHA-treated3T3-L1 adipocytes was effective in reducing monocyte migration;whereas the other fatty acids did not alter this effect compared tocontrol cells. Second, in line with the previous findings, we haveshown that both EPA and DHA are effective in increasing thesecretion of adiponectin, an anti-inflammatory adipokine, and thatDHA is more potent than EPA in inducing adiponectin secretion[34–36]. Third, the activation of pro-inflammatory signaling path-ways, in particular, ERK1/2 and p38 MAPK, is mitigated by DHAand not EPA. As mentioned, existing literature suggests that thesetwo fatty acids have differential effects in modulating the inflam-matory response depending on the cell types [14–16]. As foradipocytes, our data indicate that DHA may be more potent thanEPA in attenuating the inflammatory processes.

The mechanism(s) by which EPA and DHA exert differentbiological effects is a subject of extensive investigation. Existing

Fig. 4. Effects of n-3 fatty acids on the activation of MAPKs. Western blot analysis was carried out for the phosphorylated and total forms of ERK1/2, p38 MAPK, and JNK (A-D).GAPDH was used as an internal control for equal loading and, a representative band from each group is shown Values are expressed as mean7SEM of 9 samples in each group.ERK, extracellular signal related kinase; JNK, c-Jun NH2-terminal kinase.

Fig. 5. Effects of EPA versus DHA treatment on the incorporation of n-3 fatty acidsin phospholipids. The phospholipid fatty acid composition was determined by gaschromatography. The levels of very long chain n-3 fatty acids such as EPA, DPA, andDHA are shown. Values are mean7SEM of 3–6 samples per group.

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evidence suggests that the difference in the ability of these twofatty acids in altering membrane fluidity is one mechanism bywhich they exert differential biological effects [37–39]. Alterationin membrane fatty acid composition determines the degree ofmembrane fluidity. In our study, analysis of fatty acid compositionin phospholipids revealed distinct n-3 fatty acid profile in EPAversus DHA treated cells. As expected, both EPA and DHA treat-ments potently increased the accumulation of respective fattyacids in phospholipids. The main difference lies in the fact that EPAtreatment not only increased EPA but also DPA, its elongationproduct. It is well-known that EPA can undergo elongation viaELOVL5 to form DPA (reviewed in [18]) and this could be why wesee an increase in DPA content upon EPA treatment in adipocytes.Therefore, it seems reasonable to speculate that conversion of EPAto DPA may diminish the effect of EPA in inducing adipogenesis.This notion is supported by our data that adding DPA potentlyinhibited DHA-induced adipogenesis.

Our fatty acid analysis revealed that DHA treatment, in additionto increasing DHA, also increased EPA compared to controls. DHAcan be retroconverted to EPA via partial peroxisomal β-oxidation[40,41]. Of note, DHA supplementation reduced DPA content in

differentiated adipocytes. It has been previously reported thatDHA reduces DPA in platelet phospholipids of human subjectslikely due to reduced elongation of EPA to DPA upon DHAtreatment [42,43]. Our data are in agreement with these biochem-ical pathways involved in the intracellular processing of very longchain n-3 fatty acids and provide novel evidence that suchconversions significantly alter the n-3 fatty acids composition inphospholipids of adipocytes. Overall, our data suggest that thedifference in the magnitude of differentiation induced by thesetwo fatty acids may be due at least in part, to their differentintracellular processing which, in turn, could alter the types of n-3fatty acids accumulated in phospholipids.

However, the differential response to EPA and DHA may also bedue to a number of other factors. For example, the differentialregulation of nuclear receptors, in particular PPARγ, may cause thedivergent effects of EPA versus DHA. PPARγ is a master regulator ofadipogenesis and EPA is a potent agonist of PPARγ [44,45]. In thecurrent study, both EPA and DHA upregulated PPARγ mRNAexpression in 3T3-L1 adipocytes however, DHA was more potentin mediating this effect. Moreover, we noted that the expression ofaP2, a downstream target of PPARγ, is increased only in DHA but

Fig. 6. Effects of DPA on the proadipogenic effects of DHA. 3T3-L1 cells were pretreated and differentiated in the presence of DHA or DPA at 50 mM. In some groups, DHAwasreplaced by DPA at 10 or 25 mM concentration. After the differentiation process, the cells were stained with oil red O. The lipid droplets were dissolved in 60% isopropanoland the color intensity was measured at 490 nm in a spectrophotometer (A). RNA samples were analyzed by real time PCR for the expression of genes involved in thedifferentiation of 3T3-L1 preadipocytes (B–D). Values are expressed as mean7SEM of 6 samples in each group. UD, undifferentiated control; Diff, differentiated control;DHA, docosahexaenoic acid; DPA, docosapentaenoic acid; DHA 40þDPA 10 (DHA at 40 mMþDPA at 10 mM); and DHA 25þDPA 25 (DHA at 25 mMþDPA at 25 mM).

G. Murali et al. / Prostaglandins, Leukotrienes and Essential Fatty Acids 90 (2014) 13–21 19

not EPA treated cells compared to differentiated control cells. Thedifferential effects of EPA and DHA on PPARγ activation may alsobe due to their metabolites. Notably, a number of oxidationproducts of DHA are more potent activators of PPARγ than DHAitself [46,47] and therefore, the role of DHA oxidation products inpromoting preadipocyte differentiation cannot be ruled out.Finally, GPR120 plays an important role in adipogenesis [48] andDHA has been shown to be a potent ligand for this receptor [49].Thus, the role of GPR120 in causing the differential effects of EPAand DHA in mediating preadipocyte differentiation remainspossible.

Although fish oil is the most widely used dietary supplement inclinical settings, the individual effects of EPA versus DHA are stillnot fully understood [50]. Evidence suggests that they both havedistinct biological effects with regard to modulating lipid andglucose metabolism. For example, DHA is more beneficial than EPAin reducing cardiovascular risk factors in humans [51,52]. How-ever, EPA is also known to exert potent lipid-lowering effects inhumans [53]. A study in type 2 diabetic patients showed that DHAbut not EPA reduced the ratio of glucose to insulin in thesepatients [42]. A better knowledge on the individual effects ofEPA and DHA is needed to exploit the benefits of these fatty acidsto improve metabolic disorders in clinical settings. Regardingpreadipocyte differentiation, our data provide evidence that DHAis more potent than EPA in promoting the differentiation process.Given the possibility that DHA can induce preadipocyte differen-tiation, one would assume that DHA will promote obesity byincreasing adiposity. However, the extent of adiposity in vivo ismainly dependent on the availability of excess free fatty acids forstorage. The liver plays a central role in fatty acid metabolism andn-3 fatty acids including EPA and DHA are known to promote fattyacid oxidation and reduce fatty acid synthesis in liver [7]. There-fore, it is likely that DHA would improve AT dysfunction withoutcausing an increase in hyperplastic obesity. In fact, a number ofanimal studies have shown a reduction or no change in adiposityupon DHA supplementation [54–56]. AT dysfunction characterizedby increased AT lipolysis, secretion of pro-inflammatory cytokinesand chemokines, and accumulation of immune cells, in particular,macrophages, plays an important role in mediating the patholo-gical consequences of obesity. Therefore, our findings are relevantto considering the use of DHA supplements to ameliorate ATdysfunction in obesity which, in turn, may lead to improvementsin metabolic disorders including insulin resistance, dyslipidemia,and hepatic steatosis.

Disclosures

Ganesan Murali—No conflict of interest.Cyrus V. Desouza—No conflict of interest.Michelle E. Clevenger—No conflict of interest.Ramesh Ramalingam—No conflict of interest.Viswanathan Saraswathi—No conflict of interest.

Acknowledgments

This project was supported in part, by the American HeartAssociation Scientist Development Grant (0930335N) toV. Saraswathi. The work was conducted using the facilitiesand resources at the VA Nebraska Western Iowa Health CareSystem, Omaha. Fatty acid analysis was performed at the LipidCore Laboratory of the Mouse Metabolic Phenotyping Center atVanderbilt University (DK59637).

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