Open Access

Suppressive actions of eicosapentaenoic acid on lipid droplet formation in 3T3-L1 adipocytes

  • Elizabeth Manickam1,
  • Andrew J Sinclair1, 2 and
  • David Cameron-Smith1Email author
Lipids in Health and Disease20109:57

https://doi.org/10.1186/1476-511X-9-57

Received: 15 April 2010

Accepted: 4 June 2010

Published: 4 June 2010

Abstract

Background

Lipid droplet (LD) formation and size regulation reflects both lipid influx and efflux, and is central in the regulation of adipocyte metabolism, including adipokine secretion. The length and degree of dietary fatty acid (FA) unsaturation is implicated in LD formation and regulation in adipocytes. The aims of this study were to establish the impact of eicosapentaenoic acid (EPA; C20:5n-3) in comparison to SFA (STA; stearic acid, C18:0) and MUFA (OLA; oleic acid, C18:1n-9) on 3T3-L1 adipocyte LD formation, regulation of genes central to LD function and adipokine responsiveness. Cells were supplemented with 100 μM FA during 7-day differentiation.

Results

EPA markedly reduced LD size and total lipid accumulation, suppressing PPARγ, Cidea and D9D/SCD1 genes, distinct from other treatments. These changes were independent of alterations of lipolytic genes, as both EPA and STA similarly elevated LPL and HSL gene expressions. In response to acute lipopolysaccharide exposure, EPA-differentiated adipocytes had distinct improvement in inflammatory response shown by reduction in monocyte chemoattractant protein-1 and interleukin-6 and elevation in adiponectin and leptin gene expressions.

Conclusions

This study demonstrates that EPA differentially modulates adipogenesis and lipid accumulation to suppress LD formation and size. This may be due to suppressed gene expression of key proteins closely associated with LD function. Further analysis is required to determine if EPA exerts a similar influence on LD formation and regulation in-vivo.

Background

Adipose tissue is a complex dynamic tissue facilitating energy storage, which also exerts considerable influence on whole body metabolic function through secreted hormones and adipokines [1]. Importantly, the size of adipocytes is an important determinant of the profile of the adipokines secreted, with large adipocytes predominately releasing pro-inflammatory factors such as monocyte chemoattractant protein-1 (MCP-1) and interleukin-6 (IL-6), and reduced anti-inflammatory adipokines including leptin and adiponectin [24]. In adipocytes, triacylglycerols (TAG) are sequestered within lipid droplets (LDs), with over 95% of each mature adipocyte is composed of TAG. LD formation is the major pathway regulating adipocyte size [5].

LDs are far from inert vesicles, and are composed of a central core of neutral lipids surrounded by a phospholipid monolayer that occurs in close association with a complex array of proteins [6]. Crucial in the genesis of LDs in adipocyte differentiation from precursor fibroblasts is the developmental program initiated and regulated by key adipogenic transcription factors such as peroxisome proliferator-activated receptor γ (PPARγ) [7].

The maintenance of LDs in mature adipocytes is regulated in part both by the TAG influx dictated by enzymes including lipoprotein lipase (LPL), and the predominant enzymes regulating TAG efflux, including adipose triglyceride lipase (ATGL, also known as desnutrin and Pnpla2) and hormone sensitive lipase (HSL, also known as Lipe) [810]. The activity of these efflux lipolytic enzymes is orchestrated by protein-protein interactions with Perilipin A, a lipid droplet scaffold protein [11]. Alterations in HSL expression has limited impact on LD size, however, alterations in ATGL activity profoundly influence LD size, independent from Perilipin A activity [12]. However, Perilipin A null mice have impaired LD formation, demonstrating that Perilipin A also uniquely influences LD function [13]. Whilst the abundance of ATGL and Perilipin A demonstrate the importance of lipolytic control in regulating LD size, RNAi screening unexpectedly highlighted a close association and importance of the cell death-inducing DFF45-like effector (CIDE) domain containing protein (Cidea) with LD size [14]. The mice Cidea (homologous to humans CIDEA), previously described as a mitochondria-associating protein, associates with LDs and negatively regulates lipolysis, promoting increased LD size. Both Perilipin A and Cidea are transcriptionally regulated by PPARγ, demonstrating the importance of this pathway in the regulation of LD formation and size [15, 16].

Depending on the length and degree of unsaturation, FAs have been predicted to influence PPARγ-regulated gene expression and subsequent LD formation during adipocyte differentiation [17, 18]. Recently, adipocyte size has been noted to be influenced by the FA composition in the LDs that is regulated by delta-9 desaturase/stearoyl Co-A desaturase 1 (D9D/SCD1) gene [19] that is PPARγ-dependent [20]. Potent in the regulation of PPARγ is the long chain n-3 PUFA, eicosapentaeneoic acid (EPA). Diets enriched in EPA lower adipose tissue mass and suppress obesity development in rats [21]. Within adipocytes EPA is known to induce expression of genes for mitochondrial biogenesis and oxidative metabolism, increasing the catabolism of lipids [22, 23]. Furthermore, EPA has been previously reported to attenuate pro-inflammation in favour of anti-inflammatory adipokines [23]. Yet in spite of this, the impact of EPA on the molecular mechanisms governing the key pathways dictating adipocyte size and the biogenesis of intracellular LDs has yet to be analysed.

In this study, the differential effects of 7-day period of EPA (C20:5n-3) compared with stearic (STA, C18:0) and oleic acids (OLA, C18:1n-9) in differentiating 3T3-L1 adipocytes was assessed on the basis of the molecular factors critical in governing LD biogenesis. The ability of the resultant adipocytes to modulate the expression of pro-inflammatory, compared to anti-inflammatory adipokines upon bacterial endotoxin lipopolysaccharide (LPS) challenge were also evaluated. It was hypothesised that EPA would suppress LD formation and enhance the anti-inflammatory adipokines in response to LPS challenge.

Results

The effect of FA on LD formation in 3T3-L1 adipocytes

A representative image of LD size and number of 3T3-L1 adipocytes which underwent differentiation in the presence of differing FA is shown in Figure 1A. Adipocytes differentiated in the absence of added FA (CTRL), or with STA and OLA after 7 days resulted in larger LDs than that observed with EPA treatment. The smaller LD observed with EPA treatment reduced total lipid accumulation by 20% (P < 0.05), as measured using Oil Red O staining, but no difference was observed in other groups compared with CTRL (Figure 1B).
Figure 1

Effects of FAs on adipocyte morphology and lipid accumulation. The 3T3-L1 pre-adipocytes were incubated in growth medium supplemented with adipogenic factors (520 μM IBMX, 200 nM Dex and 2 μg/mL insulin). FAs (100 μM) of EPA, STA or OLA were included throughout 7 days of adipocyte differentiation. A) Cell morphology of differentiated adipocytes analysed with phase contrast microscopy before (a - d) and after (e - h) Oil Red O stain. B) Stained lipid fraction was measured spectrophotometrically on Day 8 at A520 nm. Values were calculated as percent lipid content versus vehicle control cells treated with 2% BSA + 100% ethanol alone (CTRL) and are expressed as means ± SEM (n = 6). Values with different superscript letters are significantly different at P < 0.05 by Tukey's post hoc test.

Gene expression of adipogenic and LD factors

The impact of EPA, in comparison to STA and OLA, on PPARγ and GLUT4, is shown in Figure 2A. EPA treatment during adipocyte differentiation decreased PPARγ by nearly 50% compared with STA or OLA. GLUT4 gene expression was unaltered by all treatments. Furthermore, genes central to LD formation (Cidea and Perilipin A) were also measured (Figure 2B). EPA significantly decreased Cidea gene expression by 50% (P < 0.05) compared with CTRL or STA. Comparatively, OLA increased Cidea gene expression. Perilipin A gene expression was unaltered from CTRL in all groups.
Figure 2

Effects of FAs on mRNA expression of (A) adipogenesis; PPARγ and GLUT4, and (B) LD size; Cidea and Perilipin A. Cells were differentiated in the presence of 100 μM FA or vehicle control. EPA, in contrast to STA and OLA, resulted in a significant inhibition of Cidea and PPARγ gene expressions but not Perilipin A or GLUT4. All values are means ± SEM of percent gene expression relative to CTRL, where cDNA levels were normalised by cyclophilin levels (n = 4). Values with different superscript letters are significantly different at P < 0.05 by Tukey's post hoc test.

Gene expression of lipolytic enzymes

As shown in Figure 3, there was a significant increased expression of LPL and HSL genes in adipocytes differentiated with all types of FAs relative to CTRL (P < 0.05). More than 2-fold increase in LPL gene expression was noted in adipocytes differentiated in the presence of EPA and STA, compared with CTRL. The expression of HSL gene was also significantly elevated by both EPA and STA, compared with CTRL (P < 0.05).
Figure 3

Effects of FAs on mRNA expression of lipolytic enzymes, LPL and HSL. Adipocytes were differentiated in the presence of 100 μM FA or vehicle control. All FA treatments, EPA, STA and OLA, increased gene expression of LPL and HSL, compared with vehicle control (CTRL). All values are means ± SEM of percent gene expression relative to CTRL, where cDNA levels were normalised by cyclophilin levels (n = 4). Values with different superscript letters are significantly different at P < 0.05 by Tukey's post hoc test.

Gene expression of D9D/SCD1

A significant elevation of D9D/SCD1 gene expression was evident in adipocyte differentiated with STA (131%) relative to CTRL (P < 0.05) (Figure 4). When EPA was introduced, the adipocytes expressed 62% less D9D/SCD1 gene expression compared with CTRL (P < 0.05).
Figure 4

Effects of FAs on mRNA expression of D9D/SCD1. Cells were differentiated in the presence of 100 μM FA or vehicle control. EPA, in contrast to STA and OLA, resulted in a significant inhibition of D9D/SCD1 gene expression. All values are means ± SEM of percent gene expression relative to CTRL, where cDNA levels were normalised by cyclophilin levels (n = 4). Values with different superscript letters are significantly different at P < 0.05 by Tukey's post hoc test.

Gene expression of inflammatory adipokines upon LPS challenge

The changes in the inflammatory adipokines upon LPS challenge in differentiated-adipocytes are shown in Table 1. The elevation of MCP-1 gene expression after LPS treatment was significantly lower in EPA-differentiated adipocytes, compared with CTRL or STA (5.6-fold vs > 13-fold) (P < 0.05). Similarly, the expression of IL-6 gene was 4.1-fold less after EPA treatment compared with STA. Alternatively, the gene expressions of adiponectin (2.2-fold vs 1.1-fold) and leptin (2.8-fold vs 1.2-fold) were significantly elevated upon LPS challenge in EPA compared with STA treatment.
Table 1

Gene expression of inflammatory adipokines upon LPS challenge in adipocytes after differentiation with EPA, STA, OLA or vehicle control

Gene expression of adipokines (fold change/unstimulated)

FA treatment (100 μM)

 

CTRL

EPA

STA

OLA

MCP-1

13.9 ± 3.1a

5.6 ± 0.8b

13.4 ± 3.3a

6.6 ± 1.8ab

IL-6

7.2 ± 0.8ab

5.5 ± 0.8b

9.6 ± 0.9a

5.2 ± 0.8b

Adiponectin

1.4 ± 0.1ab

2.0 ± 0.4b

1.1 ± 0.2a

1.3 ± 0.2ab

Leptin

0.9 ± 0.2a

2.8 ± 1.0b

1.2 ± 0.2a

1.0 ± 0.3a

Values are means ± SEM of n = 4, with different superscript letters are significantly different at P < 0.05 by Tukey's post hoc test.

Discussion

LDs are important organelles, not only for cellular energy storage, but emerging research demonstrates a central role in intracellular signalling and adipokine regulation. Thus, the formation and regulation of LDs may contribute to the metabolic disease progression towards type 2 diabetes and cardiovascular disease evident in obesity. FAs are important in both providing the substrate for LD formation, and may also be central in regulating LD formation and size. Previously, long chain n-3 PUFA have been reported to suppress LD formation [24, 25], although comparison to other FA and putative mechanisms is lacking. The present study demonstrated a marked suppression of LD formation in 3T3-L1 adipocytes maintained in the presence of EPA, when compared to either a SFA or MUFA. Of the mechanisms governing LD formation and regulation, EPA suppressed PPARγ, Cidea and D9D/SCD1 gene expressions, while maintaining the expression of lipolytic genes, including LPL and HSL. A key regulator of adipogenesis is PPARγ, and FA sensitive transcriptional regulator of many thousands of adipocyte-specific genes [7].

Specifically, both n-6 and n-3 PUFA species directly influence PPAR activity [23]. Although known as an acute agonist and regulator of the PPARγ gene [26], chronic EPA exposure was shown to suppress this gene expression. Many of the subsequent genes analysed in the present study are PPAR-dependent, yet the responsiveness was varied, suggesting alteration of PPAR activity was not the sole regulating factor suppressing LD formation in the presence of EPA.

Cidea and Perilipin A are proteins highly localised to LDs of adipose tissue that are important for FA esterification to TAG and lipid mobilisation [14, 27]. Cidea and Perilipin A are known to be closely regulated by PPARγ [16]. Cidea acts as a shield for fat storage that drives lipid accumulation even in cells that rarely store fat [28]. Similarly, Cidea expression is abundantly upregulated with high SFA feeding [14]. Conversely, animals or humans with Cidea deficiency display lean phenotypes with resistance to diet-induced obesity [29]. In this study, Cidea gene expression was downregulated in adipocytes treated with EPA but not after STA or OLA treatments.

Perilipins are a class of proteins closely associated with LD formation in adipocytes [30], yet the critical relevance of this gene in the regulation of lipid mobilisation upon nutritional challenges in adipocytes is uncertain. An elevated Perilipin A level in obese humans [31] is inconsistent with decreased diet-induced obesity in Perilipin A-overexpressed mice [32]. In this study, we found that EPA significantly affected Cidea but not Perilipin A. The substantial reduction in Cidea gene expression by EPA, but not by STA or OLA, is consistent with the decreased LD size and PPARγ, suggesting that Cidea (along with PPARγ) is the prime target for EPA in adipocytes, not Perilipin A.

Another possible mechanism governing LD formation is linked to D9D/SCD1. Desaturases are other enzymes that are greatly affected by the degree of unsaturation of FAs. D9D/SCD1 is one particular desaturase that has been highly discussed in terms of SFA-rich diet-induced obesity. D9D/SCD1 is a key regulatory enzyme required to convert SFA to MUFA that participate in lipid metabolism in adipocytes [33, 34]. The importance of D9D/SCD1 expression in the formation of larger adipocytes has also been recently noted [19]. Contrary to the upregulation shown by SFA and MUFA, dietary PUFA (arachidonic and α-linolenic acids) consistently decreased the expression of D9D/SCD1 in liver and adipose tissue of obese models [35, 36]. Similar to the shorter chain n-3 PUFA α-linolenic acids, our study is the first to confirm a distinct downregulation of D9D/SCD1 in adipocytes after longer chain n-3 PUFA EPA treatment, in contrast to the upregulation shown by STA and OLA.

Like many other adipocyte-specific genes, D9D/SCD1 is also upregulated by PPARγ during adipogenesis, while PPARγ expression is not controlled by D9D/SCD1 [37]. Whilst D9D/SCD1 is positively correlated with PPARγ to increase lipogenesis in adipocytes [20, 37], the significance of reducing the D9D/SCD1 expression to reverse obesity has been constantly discussed [38, 39]. Many anti-obesity compounds are designed with the aim of reducing the level of D9D/SCD1 [40, 41], similar to the effect shown here by EPA. Bigger LDs has been shown to strongly correlate with the upregulation of PPARγ, D9D/SCD1 and Cidea in adipocytes supplemented with STA and OLA. As illustrated in Figure 5, it is clear that the anti-adipogenic effect of EPA is regulated by the reduction in PPARγ level, which decreased D9D/SCD1 and Cidea gene expressions, followed by a decline in lipid accumulation to induce smaller LDs. However, it is not known whether D9D/SCD1 and Cidea are independent of each other.
Figure 5

Possible mechanism involved in FA-regulated LD formation in adipocytes. Both SFA (stearic acid) and n-3 PUFA (eicosapentaenoic acid) have similar magnitude of effects towards lipolysis controlled by LPL and HSL. In contrast to upregulation shown by SFA, n-3 PUFA decreased PPARγ level, which may be responsible for the reduction of D9D/SCD1 and Cidea expressions, leading to smaller LDs. The levels of GLUT4 and Perilipin A remain unchanged upon FA treatment.

It is plausible that modifications in lipolyic pathways influence LD formation and size regulation. LPL and HSL are the major rate-limiting lipases for a balanced lipolysis and lipogenesis in adipocytes. In this study the degree of unsaturation of FAs did not alter the mRNA levels of LPL and HSL in adipocytes. Both EPA and STA (and OLA to some extent) affect the expressions of these lipases at the same magnitude in adipocytes. This results support previous studies on the influence of FAs on lipolytic enzymes gene expressions. Dietary fish oil supplementation increased LPL gene expression in adipose tissue of humans and rats [42, 43]. In comparison to other FAs, n-3 PUFA-rich perilla oil did not change LPL expression in rat adipose tissue although a reduced adipogenesis was observed [44]. Similarly, HSL level is only slightly affected by the degree of FA unsaturation, confirming a previous notion [45]. It is speculated that like other FAs, EPA may play a role in lipolytic regulation, however this does not influence the biogenesis of LDs in adipocytes.

Adipocytes produce numerous amounts of inflammatory adipokines upon stimulation. Hypertrophied adipocytes are insulin resistant, and demonstrate greater secretion of low grade inflammatory cytokines, chemokines and vascular proteins [46]. Our study demonstrated that adipocytes differentiated with EPA suppressed gene expressions of LPS-induced pro-inflammatory adipokines, MCP-1 and IL-6. Alternatively, similar treatment resulted in elevated adiponectin and leptin gene expressions compared with SFA, complementing data described elsewhere [47, 48]. Thus, EPA-treated adipocytes, with smaller LDs, have improved inflammatory response.

Conclusion

EPA, being the predominant long chain n-3 PUFA found in marine oils, suppresses LD formation and size in 3T3-L1 adipocytes, in comparison to other FAs. Several lines of evidence suggest a beneficial impact of EPA on lipid accumulation and body weight regulation [21, 23, 49]. Suppression of LD formation and size may be one novel contributing factor. In addition to examining LD size, we explored the gene analysis of known regulators of LD function. Interestingly, addition of EPA during adipocyte differentiation decreased PPARγ, Cidea and D9D/SCD1 without affecting GLUT4 and Perilipin A expressions, as compared with SFA. Such effects concomitantly suppressed the total lipid accumulation and biogenesis of LDs. Complementing this data is altered expression of adipokines following acute LPS exposure, with EPA-treated adipocytes expressing improved inflammatory response. Thus, EPA is a novel regulator of LD formation, a key intracellular organelle central to the function and regulation of adipocyte metabolism.

Methods

Reagents

Dexamethasone (Dex; D4902), isobutylmethylxanthine (IBMX; I5879), insulin (I5500), low-endotoxin FA-free BSA (A8806), LPS (L6529), EPA (44864), STA (85679) and OLA (O1383) (all FAs were with purity > 99%) were obtained from Sigma (St. Louis, MO). Fetal bovine serum (FBS) and DMEM were purchased from Invitrogen (Carlsbad, CA). FAs were dissolved in 100% ethanol as stock solutions of 100 mM while LPS was dissolved in dH2O at 1 mg/mL stock. All stock solutions were stored in the dark at -20°C before use. All other reagents were of analytical grade.

Cell culture

The 3T3-L1 mouse embryo fibroblasts were obtained from the American Type Culture Collection (Rockville, MD). The adipocyte cell culture and their differentiation from 3T3-L1 pre-adipocytes were carried out as described previously [50]. Briefly, cells were seeded in 6-, 12- and 24-well tissue culture plates and maintained in growth medium containing DMEM supplemented with 10% FBS, 2 mM/L glutamine, 100 U/L penicillin and 100 μg/mL streptomycin in a humidified atmosphere of 95% air/5% CO2 at 37°C. At post-confluent, adipogenesis of 3T3-L1 was induced with growth media containing 520 μM IBMX, 200 nM Dex and 2 μg/mL insulin. After 3 days of exposure to the differentiation medium, cells were maintained in growth medium containing insulin alone. FA treatment was introduced throughout all 7 days of cell differentiation at physiological concentration of 100 μM. STA was diluted in warm growth medium containing 2% BSA and incubated for 4 hours at 37°C until dissolved. All other FAs were freshly prepared from the stock solution and diluted with growth medium containing 2% BSA. A corresponding amount of 2% BSA + 100% ethanol was used as the vehicle control (CTRL). For LPS stimulation, mature adipocytes were serum starved for 2 h prior to treatment with 1 μg/mL of LPS for 60 min at 37°C to induce inflammation.

Oil Red O staining

After appropriate treatment, adipocytes were washed with cold phosphate-buffered saline (PBS; pH 7.4) and fixed with 4% paraformaldehyde solution in PBS. Cells were stained with freshly prepared Oil Red O dye (0.5% (w/v) dissolved in isopropanol and diluted at 3:2 ratio of dye:water). Cells were washed thoroughly with distilled water prior to quantification. Spectrophotometric analysis of the stain was performed by dissolving the stained LDs with isopropanol and measuring at A520 nm [51]. The values were calculated as percentage of the vehicle control and expressed as means ± SEM (n = 6).

RNA isolation and gene expression quantification

Total RNA was extracted from 3T3-L1 adipocytes using Tri-reagent (PE Applied Biosystems, Foster City, CA). The purity of the RNA was measured at A260 nm/A280 nm using Nanodrop 1000 (Thermo Fisher Scientific Inc, MA, USA) with ratio values of ~2.0. The cDNAs was synthesised from 2 μg of RNA using high capacity RNA-to-cDNA reverse transcriptase kit in a total of 20 μL of reaction volume (PE Applied Biosystems). Reverse transcription was performed with sample incubation at 42°C for 1 h, followed by 99°C for 5 min and 4°C for another 5 min. RT-PCR amplifications were performed from 1 μL cDNA diluted at 1:20 using each of the gene-specific primer sets (GeneWorks Pty Ltd, SA, Australia). The oligonucleotide sequence of the forward (sense) and reverse (antisense) primers used for amplification were as in Table 2. Each primer set was used at a concentration of 200 μM in a final volume of 14 μL using Power SYBR® Green PCR Master Mix (PE Applied Biosystems). All PCR reactions were performed using a fluorometric thermal cycler (7500 Fast Real-Time PCR System, PE Applied Biosystems). The ΔΔCT method was used to measure relative quantification, where values were normalised to the reference gene (cyclophilin). Individual CT values are means of triplicate measurements, with repeatability measurement of < 0.5. Separate control experiments were used to demonstrate that the efficiency of target and reference amplifications were equal.
Table 2

Gene specific forward and reverse primer sequences used for RT-PCR

Gene

Forward primer (5' - 3')

Reverse primer (5' - 3')

NCBI accession number

Adiponectin

AAGGACAAGGCCGTTCTCT

TATGGGTAGTTGCAGTCAGTTGG

NM_009605

Cidea

TGCTCTTCTGTATCGCCCAGT

GCCGTGTTAAGGAATCTGCTG

NM_007702

Cyclophilin

TCTGCTGTCTTTGGAACTTTGTC

CTGATGGCGAGCCCTTG

NM_008907

D9D/SCD1

ATGCCGGCCCACATGCTCCAA

TCAGCTACTCTTGTGACTCCC

NM_009127

GLUT4

TGTGGCCTTCTTTGAGATTGG

CTGAAGAGCTCGGCCCACAA

NM_009204

HSL

CCTACTGCTGGGCTGTCAA

CCATCTCGCACCCTCACT

NM_010719

IL-6

ACAAGTCGGAGGCTTAATTAGACAT

TTGCCATTGCACAACTCTATTC

NM_031168

Leptin

TCCAGAAAGTCCAGGATGACAC

CACATTTTGGGAAGGCAGG

NM_008493

LPL

AGTAGACTGGTTGTATCGGG

AGCGTCATCAGGAGAAAGG

NM_008509

MCP-1

CTTCCTCCACCACCATGCA

CCAGCCGGCAACTGTGA

NM_011333

Perilipin A

TGCTGGATGGAGACCTC

ACCGGCTCCATGCTCCA

NM_001113471

PPARγ

GGAATGGGAGTGGTCATCCA

CCCACCAACTTCGGAATC

NM_001127330

Statistical analysis

All results are expressed as means ± SEM. The statistical significance of differences among groups was determined by one-way analysis of variance (ANOVA) using the SPSS package program version 17.0 (SPSS, Chicago, IL). The results were considered to be significant if the value of P was < 0.05 using Tukey's post hoc test.

Abbreviations

ATGL

adipose triglyceride lipase or desnutrin or Pnpla2

Cidea

cell death-inducing DNA fragmentation factor alpha subunit-like effector A

D9D/SCD1

delta-9 desaturase/stearoyl Co-A desaturase 1

Dex

dexamethasone

EPA

eicosapentaenoic acid (C20:5n-3)

FA

fatty acid

GLUT4

glucose transporter type 4

HSL

hormone sensitive lipase or Lipe

IBMX

isobutylmethylxanthine

IL-6

interleukin-6

LD

lipid droplet

LPL

lipoprotein lipase

LPS

lipopolysaccharide

MCP-1

monocyte chemoattractant protein-1

OLA

oleic acid (C18:1n-9)

PPARγ

peroxisome proliferator-activated receptor gamma

SFA

saturated fatty acids

STA

stearic acid (C18:0)

TAG

triacylglycerols or triglycerides

Declarations

Acknowledgements

The authors wish to thank for the financial support through Alfred Deakin Postdoctoral Fellowship program awarded to EM.

Authors’ Affiliations

(1)
Molecular Nutrition Unit, School of Exercise and Nutrition Sciences, Faculty of Health, Medicine, Nursing and Behavioural Sciences, Deakin University
(2)
School of Medicine, Faculty of Health, Medicine, Nursing and Behavioural Sciences, Deakin University

References

  1. Wozniak SE, Gee LL, Wachtel MS, Frezza EE: Adipose tissue: the new endocrine organ? A review article. Digestive diseases and sciences. 2009, 54 (9): 1847-1856. 10.1007/s10620-008-0585-3View ArticlePubMedGoogle Scholar
  2. McLaughlin T, Deng A, Gonzales O, Aillaud M, Yee G, Lamendola C, Abbasi F, Connolly AJ, Sherman A, Cushman SW: Insulin resistance is associated with a modest increase in inflammation in subcutaneous adipose tissue of moderately obese women. Diabetologia. 2008, 51 (12): 2303-2308. 10.1007/s00125-008-1148-zPubMed CentralView ArticlePubMedGoogle Scholar
  3. Koska J, Stefan N, Dubois S, Trinidad C, Considine RV, Funahashi T, Bunt JC, Ravussin E, Permana PA: mRNA concentrations of MIF in subcutaneous abdominal adipose cells are associated with adipocyte size and insulin action. International journal of obesity (2005). 2009, 33 (8): 842-850. 10.1038/ijo.2009.106View ArticleGoogle Scholar
  4. Tejero ME, Proffitt JM, Rodriguez IP, Hubbard G, Freeland-Graves JH, Peebles KW, Cole SA, Comuzzie A: Adipokine expression is associated with adipocyte volume in baboons. Cytokine. 2008, 41 (2): 150-154. 10.1016/j.cyto.2007.11.005View ArticlePubMedGoogle Scholar
  5. Murphy S, Martin S, Parton RG: Lipid droplet-organelle interactions; sharing the fats. Biochimica et biophysica acta. 2009, 1791 (6): 441-447.View ArticlePubMedGoogle Scholar
  6. Farese RV, Walther TC: Lipid droplets finally get a little R-E-S-P-E-C-T. Cell. 2009, 139 (5): 855-860. 10.1016/j.cell.2009.11.005PubMed CentralView ArticlePubMedGoogle Scholar
  7. Schoonjans K, Staels B, Auwerx J: The peroxisome proliferator activated receptors (PPARS) and their effects on lipid metabolism and adipocyte differentiation. Biochimica et biophysica acta. 1996, 1302 (2): 93-109.View ArticlePubMedGoogle Scholar
  8. Lafontan M, Langin D: Lipolysis and lipid mobilization in human adipose tissue. Progress in lipid research. 2009, 48 (5): 275-297. 10.1016/j.plipres.2009.05.001View ArticlePubMedGoogle Scholar
  9. Ahmadian M, Wang Y, Sul HS: Lipolysis in adipocytes. The international journal of biochemistry & cell biology. 2009, 42 (5): 555-559. 10.1016/j.biocel.2009.12.009View ArticleGoogle Scholar
  10. Zimmermann R, Strauss JG, Haemmerle G, Schoiswohl G, Birner-Gruenberger R, Riederer M, Lass A, Neuberger G, Eisenhaber F, Hermetter A: Fat mobilization in adipose tissue is promoted by adipose triglyceride lipase. Science (New York, NY. 2004, 306 (5700): 1383-1386.View ArticleGoogle Scholar
  11. Brasaemle DL, Rubin B, Harten IA, Gruia-Gray J, Kimmel AR, Londos C: Perilipin A increases triacylglycerol storage by decreasing the rate of triacylglycerol hydrolysis. The Journal of biological chemistry. 2000, 275 (49): 38486-38493. 10.1074/jbc.M007322200View ArticlePubMedGoogle Scholar
  12. Miyoshi H, Perfield JW, Obin MS, Greenberg AS: Adipose triglyceride lipase regulates basal lipolysis and lipid droplet size in adipocytes. Journal of cellular biochemistry. 2008, 105 (6): 1430-1436. 10.1002/jcb.21964PubMed CentralView ArticlePubMedGoogle Scholar
  13. Hickenbottom SJ, Kimmel AR, Londos C, Hurley JH: Structure of a lipid droplet protein; the PAT family member TIP47. Structure. 2004, 12 (7): 1199-1207. 10.1016/j.str.2004.04.021View ArticlePubMedGoogle Scholar
  14. Puri V, Ranjit S, Konda S, Nicoloro SM, Straubhaar J, Chawla A, Chouinard M, Lin C, Burkart A, Corvera S: Cidea is associated with lipid droplets and insulin sensitivity in humans. Proceedings of the National Academy of Sciences of the United States of America. 2008, 105 (22): 7833-7838. 10.1073/pnas.0802063105PubMed CentralView ArticlePubMedGoogle Scholar
  15. Arimura N, Horiba T, Imagawa M, Shimizu M, Sato R: The peroxisome proliferator-activated receptor gamma regulates expression of the perilipin gene in adipocytes. The Journal of biological chemistry. 2004, 279 (11): 10070-10076. 10.1074/jbc.M308522200View ArticlePubMedGoogle Scholar
  16. Viswakarma N, Yu S, Naik S, Kashireddy P, Matsumoto K, Sarkar J, Surapureddi S, Jia Y, Rao MS, Reddy JK: Transcriptional regulation of Cidea, mitochondrial cell death-inducing DNA fragmentation factor alpha-like effector A, in mouse liver by peroxisome proliferator-activated receptor alpha and gamma. The Journal of biological chemistry. 2007, 282 (25): 18613-18624. 10.1074/jbc.M701983200View ArticlePubMedGoogle Scholar
  17. Bernlohr DA, Coe NR, Simpson MA, Hertzel AV: Regulation of gene expression in adipose cells by polyunsaturated fatty acids. Advances in experimental medicine and biology. 1997, 422: 145-156.View ArticlePubMedGoogle Scholar
  18. Takahashi Y, Ide T: Effect of dietary fats differing in degree of unsaturation on gene expression in rat adipose tissue. Annals of nutrition & metabolism. 1999, 43 (2): 86-97. 10.1159/000012772View ArticleGoogle Scholar
  19. Matsubara Y, Kano K, Kondo D, Mugishima H, Matsumoto T: Differences in adipocytokines and fatty acid composition between two adipocyte fractions of small and large cells in high-fat diet-induced obese mice. Annals of nutrition & metabolism. 2009, 54 (4): 258-267. 10.1159/000229506View ArticleGoogle Scholar
  20. Yao-Borengasser A, Rassouli N, Varma V, Bodles AM, Rasouli N, Unal R, Phanavanh B, Ranganathan G, McGehee RE, Kern PA: Stearoyl-coenzyme A desaturase 1 gene expression increases after pioglitazone treatment and is associated with peroxisomal proliferator-activated receptor-gamma responsiveness. The Journal of clinical endocrinology and metabolism. 2008, 93 (11): 4431-4439. 10.1210/jc.2008-0782PubMed CentralView ArticlePubMedGoogle Scholar
  21. Rustan AC, Hustvedt BE, Drevon CA: Dietary supplementation of very long-chain n-3 fatty acids decreases whole body lipid utilization in the rat. Journal of lipid research. 1993, 34 (8): 1299-1309.PubMedGoogle Scholar
  22. Flachs P, Horakova O, Brauner P, Rossmeisl M, Pecina P, Franssen-van Hal N, Ruzickova J, Sponarova J, Drahota Z, Vlcek C: Polyunsaturated fatty acids of marine origin upregulate mitochondrial biogenesis and induce beta-oxidation in white fat. Diabetologia. 2005, 48 (11): 2365-2375. 10.1007/s00125-005-1944-7View ArticlePubMedGoogle Scholar
  23. Flachs P, Rossmeisl M, Bryhn M, Kopecky J: Cellular and molecular effects of n-3 polyunsaturated fatty acids on adipose tissue biology and metabolism. Clin Sci (Lond). 2009, 116 (1): 1-16. 10.1042/CS20070456View ArticleGoogle Scholar
  24. Madsen L, Petersen RK, Kristiansen K: Regulation of adipocyte differentiation and function by polyunsaturated fatty acids. Biochimica et biophysica acta. 2005, 1740 (2): 266-286.View ArticlePubMedGoogle Scholar
  25. Lee MS, Kwun IS, Kim Y: Eicosapentaenoic acid increases lipolysis through up-regulation of the lipolytic gene expression and down-regulation of the adipogenic gene expression in 3T3-L1 adipocytes. Genes & nutrition. 2008, 2 (4): 327-330. 10.1007/s12263-007-0068-8View ArticleGoogle Scholar
  26. Chambrier C, Bastard JP, Rieusset J, Chevillotte E, Bonnefont-Rousselot D, Therond P, Hainque B, Riou JP, Laville M, Vidal H: Eicosapentaenoic acid induces mRNA expression of peroxisome proliferator-activated receptor gamma. Obes Res. 2002, 10 (6): 518-525. 10.1038/oby.2002.70View ArticlePubMedGoogle Scholar
  27. Garcia A, Subramanian V, Sekowski A, Bhattacharyya S, Love MW, Brasaemle DL: The amino and carboxyl termini of perilipin a facilitate the storage of triacylglycerols. The Journal of biological chemistry. 2004, 279 (9): 8409-8416. 10.1074/jbc.M311198200View ArticlePubMedGoogle Scholar
  28. Qi J, Gong J, Zhao T, Zhao J, Lam P, Ye J, Li JZ, Wu J, Zhou HM, Li P: Downregulation of AMP-activated protein kinase by Cidea-mediated ubiquitination and degradation in brown adipose tissue. The EMBO journal. 2008, 27 (11): 1537-1548. 10.1038/emboj.2008.92PubMed CentralView ArticlePubMedGoogle Scholar
  29. Zhou Z, Yon Toh S, Chen Z, Guo K, Ng CP, Ponniah S, Lin SC, Hong W, Li P: Cidea-deficient mice have lean phenotype and are resistant to obesity. Nature genetics. 2003, 35 (1): 49-56. 10.1038/ng1225View ArticlePubMedGoogle Scholar
  30. Greenberg AS, Egan JJ, Wek SA, Moos MC, Londos C, Kimmel AR: Isolation of cDNAs for perilipins A and B: sequence and expression of lipid droplet-associated proteins of adipocytes. Proceedings of the National Academy of Sciences of the United States of America. 1993, 90 (24): 12035-12039. 10.1073/pnas.90.24.12035PubMed CentralView ArticlePubMedGoogle Scholar
  31. Kern PA, Di Gregorio G, Lu T, Rassouli N, Ranganathan G: Perilipin expression in human adipose tissue is elevated with obesity. The Journal of clinical endocrinology and metabolism. 2004, 89 (3): 1352-1358. 10.1210/jc.2003-031388View ArticlePubMedGoogle Scholar
  32. Miyoshi H, Souza SC, Endo M, Sawada T, Perfield JW, Shimizu C, Stancheva Z, Nagai S, Strissel KJ, Yoshioka N: Perilipin overexpression in mice protects against diet-induced obesity. Journal of lipid research. 2009, 51 (5): 975-982. 10.1194/jlr.M002352View ArticlePubMedGoogle Scholar
  33. Miyazaki M, Ntambi JM: Role of stearoyl-coenzyme A desaturase in lipid metabolism. Prostaglandins, leukotrienes, and essential fatty acids. 2003, 68 (2): 113-121. 10.1016/S0952-3278(02)00261-2View ArticlePubMedGoogle Scholar
  34. Ntambi JM, Miyazaki M: Regulation of stearoyl-CoA desaturases and role in metabolism. Progress in lipid research. 2004, 43 (2): 91-104. 10.1016/S0163-7827(03)00039-0View ArticlePubMedGoogle Scholar
  35. Jones BH, Maher MA, Banz WJ, Zemel MB, Whelan J, Smith PJ, Moustaid N: Adipose tissue stearoyl-CoA desaturase mRNA is increased by obesity and decreased by polyunsaturated fatty acids. The American journal of physiology. 1996, 271 (1 Pt 1): E44-49.PubMedGoogle Scholar
  36. Sessler AM, Kaur N, Palta JP, Ntambi JM: Regulation of stearoyl-CoA desaturase 1 mRNA stability by polyunsaturated fatty acids in 3T3-L1 adipocytes. The Journal of biological chemistry. 1996, 271 (47): 29854-29858. 10.1074/jbc.271.47.29854View ArticlePubMedGoogle Scholar
  37. Christianson JL, Nicoloro S, Straubhaar J, Czech MP: Stearoyl-CoA desaturase 2 is required for peroxisome proliferator-activated receptor gamma expression and adipogenesis in cultured 3T3-L1 cells. The Journal of biological chemistry. 2008, 283 (5): 2906-2916. 10.1074/jbc.M705656200View ArticlePubMedGoogle Scholar
  38. MacDonald ML, Singaraja RR, Bissada N, Ruddle P, Watts R, Karasinska JM, Gibson WT, Fievet C, Vance JE, Staels B: Absence of stearoyl-CoA desaturase-1 ameliorates features of the metabolic syndrome in LDLR-deficient mice. Journal of lipid research. 2008, 49 (1): 217-229. 10.1194/jlr.M700478-JLR200View ArticlePubMedGoogle Scholar
  39. Ntambi JM, Miyazaki M, Stoehr JP, Lan H, Kendziorski CM, Yandell BS, Song Y, Cohen P, Friedman JM, Attie AD: Loss of stearoyl-CoA desaturase-1 function protects mice against adiposity. Proceedings of the National Academy of Sciences of the United States of America. 2002, 99 (17): 11482-11486. 10.1073/pnas.132384699PubMed CentralView ArticlePubMedGoogle Scholar
  40. Choi H, Eo H, Park K, Jin M, Park EJ, Kim SH, Park JE, Kim S: A water-soluble extract from Cucurbita moschata shows anti-obesity effects by controlling lipid metabolism in a high fat diet-induced obesity mouse model. Biochemical and biophysical research communications. 2007, 359 (3): 419-425. 10.1016/j.bbrc.2007.05.107View ArticlePubMedGoogle Scholar
  41. Li CS, Belair L, Guay J, Murgasva R, Sturkenboom W, Ramtohul YK, Zhang L, Huang Z: Thiazole analog as stearoyl-CoA desaturase 1 inhibitor. Bioorg Med Chem Lett. 2009, 19 (17): 5214-5217. 10.1016/j.bmcl.2009.07.015View ArticlePubMedGoogle Scholar
  42. Khan S, Minihane AM, Talmud PJ, Wright JW, Murphy MC, Williams CM, Griffin BA: Dietary long-chain n-3 PUFAs increase LPL gene expression in adipose tissue of subjects with an atherogenic lipoprotein phenotype. Journal of lipid research. 2002, 43 (6): 979-985.PubMedGoogle Scholar
  43. Yoshida T, Gotoda T, Okubo M, Iizuka Y, Ishibashi S, Kojima T, Murakami T, Murase T, Yamada N: A Japanese patient with lipoprotein lipase deficiency homozygous for the Gly188Glu mutation prevalent worldwide. Journal of atherosclerosis and thrombosis. 2000, 7 (1): 45-49.View ArticlePubMedGoogle Scholar
  44. Okuno M, Kajiwara K, Imai S, Kobayashi T, Honma N, Maki T, Suruga K, Goda T, Takase S, Muto Y: Perilla oil prevents the excessive growth of visceral adipose tissue in rats by down-regulating adipocyte differentiation. The Journal of nutrition. 1997, 127 (9): 1752-1757.PubMedGoogle Scholar
  45. Raclot T, Holm C, Langin D: Fatty acid specificity of hormone-sensitive lipase. Implication in the selective hydrolysis of triacylglycerols. Journal of lipid research. 2001, 42 (12): 2049-2057.PubMedGoogle Scholar
  46. Wueest S, Rapold RA, Rytka JM, Schoenle EJ, Konrad D: Basal lipolysis, not the degree of insulin resistance, differentiates large from small isolated adipocytes in high-fat fed mice. Diabetologia. 2009, 52 (3): 541-546. 10.1007/s00125-008-1223-5View ArticlePubMedGoogle Scholar
  47. Flachs P, Mohamed-Ali V, Horakova O, Rossmeisl M, Hosseinzadeh-Attar MJ, Hensler M, Ruzickova J, Kopecky J: Polyunsaturated fatty acids of marine origin induce adiponectin in mice fed a high-fat diet. Diabetologia. 2006, 49 (2): 394-397. 10.1007/s00125-005-0053-yView ArticlePubMedGoogle Scholar
  48. Itoh M, Suganami T, Satoh N, Tanimoto-Koyama K, Yuan X, Tanaka M, Kawano H, Yano T, Aoe S, Takeya M: Increased adiponectin secretion by highly purified eicosapentaenoic acid in rodent models of obesity and human obese subjects. Arteriosclerosis, thrombosis, and vascular biology. 2007, 27 (9): 1918-1925. 10.1161/ATVBAHA.106.136853View ArticlePubMedGoogle Scholar
  49. Herzberg GR, Skinner C: Differential accumulation and release of long-chain n-3 fatty acids from liver, muscle, and adipose tissue triacylglycerols. Canadian journal of physiology and pharmacology. 1997, 75 (8): 945-951. 10.1139/cjpp-75-8-945PubMedGoogle Scholar
  50. Green H, Kehinde O: An established preadipose cell line and its differentiation in culture. II. Factors affecting the adipose conversion. Cell. 1975, 5 (1): 19-27. 10.1016/0092-8674(75)90087-2View ArticlePubMedGoogle Scholar
  51. Ramirez-Zacarias JL, Castro-Munozledo F, Kuri-Harcuch W: Quantitation of adipose conversion and triglycerides by staining intracytoplasmic lipids with Oil red O. Histochemistry. 1992, 97 (6): 493-497. 10.1007/BF00316069View ArticlePubMedGoogle Scholar

Copyright

© Manickam et al; licensee BioMed Central Ltd. 2010

This article is published under license to BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Advertisement