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Journal of Animal Science - Animal Physiology

Heat stress enhances adipogenic differentiation of subcutaneous fat depot–derived porcine stromovascular cells1

 

This article in JAS

  1. Vol. 93 No. 8, p. 3832-3842
     
    Received: Mar 06, 2015
    Accepted: May 07, 2015
    Published: July 10, 2015


    2 Corresponding author(s): kajuwon@purdue.edu
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doi:10.2527/jas.2015-9074
  1. H. Qu,
  2. S. S. Donkin and
  3. K. M. Ajuwon 2
  1. Department of Animal Sciences, Purdue University, West Lafayette, IN 47907-2054

Abstract

Heat stress (HS) results from excessive heat load on animals such that all adaptive mechanisms used to dissipate the heat do not return the body to normal body temperature. In pigs, HS results in increased fat deposition compared with pair-fed animals in a thermoneutral environment. Although there is evidence that HS increases activity of lipoprotein lipase (LPL) in adipose tissue of heat stressed pigs, the fundamental causes of the increased adiposity are still unknown. It remains unclear whether HS directly alters metabolism in adipocytes. Therefore, to understand the mechanism of HS effects on porcine adipocytes, we used an in vitro adipocyte differentiation model to characterize cellular responses that occur during differentiation of pig adipocytes. Preadipocytes (stromovascular cells) were differentiated for 9 d at a normal (37°C) or HS (41.5°C) temperature under 5% CO2. Expressions of HS genes such as heat shock proteins (HSP; HSP27, HSP60, HSP70, and HSP90), adipogenic markers peroxisome proliferator-activated receptor γ (PPARγ), CCAAT/enhancer binding proteins α (C/EBPα), fatty acid synthase (FAS), adipocyte protein 2 (aP2), fatty acid translocase 36 (CD36), fatty acid transport protein 4 (FATP4), fatty acid transport protein 6 (FATP6), LPL, glucose transporter protein type 4 (GLUT4), phosphoenolpyruvate carboxykinase 1 (PCK1 or PEPCK-C), and glycerol kinase (GK) and adipokines (adiponectin and leptin) were determined by real-time-PCR and immunoblotting or ELISA. Cellular triglyceride (TAG) and ATP concentrations were also determined. As expected, HS increased (P < 0.05) the expressions of HSP genes. There was no HS treatment effect on the level of PPARγ, although C/EBPα was induced (P < 0.05) in HS. So it remains unclear whether HS affects adipocyte differentiation. However, HS leads to increased expressions of genes involved in fatty acid uptake and TAG synthesis (FAS, aP2, CD36, FATP4, FATP6, LPL, GLUT4, PCK1, and GK). This is supported by increased cellular TAG under HS. Therefore, HS promotes increased adipocyte TAG storage, perhaps through upregulation of genes involved in fatty acid uptake and TAG synthesis.



INTRODUCTION

Heat stress (HS) occurs when animals are unable to dissipate excess body heat (West, 2003). Pigs are especially vulnerable to HS because they lack functional sweat glands on their skin. Pigs exposed to HS suffer reduced performance (Spencer et al., 2005). Effects of HS in pigs include increased plasma insulin and increased N(τ)-methylhistidine but decreased plasma NEFA (Pearce et al., 2013a). Heat stress results in increased carcass fat (Bridges et al., 1998; Collin et al., 2001) in the pig when compared with pair-fed animals at a thermoneutral temperature. Although precise mechanisms of this effect are unknown, elevated plasma insulin, a lipogenic hormone, may be a contributing factor during HS (Pearce et al., 2013a). Increased insulin concentration may also be responsible for the increased expression and activity of lipoprotein lipase (LPL), another adipogenic enzyme, in the adipose tissue of heat stressed pigs (Kouba et al., 2001).

Despite the likely implication of insulin in the increased fat mass of heat stressed pigs, there are significant inconsistencies in the regulation of serum insulin levels in pigs under HS (Williams et al., 2013; Pearce et al., 2014), suggesting that additional mechanisms may be involved. Experiments are needed to elucidate the precise mechanisms of HS response in the adipose tissue of the pigs to establish implications for the increased adipose accretion in pigs under HS. Using an in vitro differentiation model of pig preadipocytes exposed to mild HS recapitulates the effects of medium- to long-term HS in pigs. Unlike acute HS, chronic HS induces more adaptive responses and nonlethal metabolic alterations (Horowitz, 2002), more akin to an in vivo adaptive response in pigs. Therefore, the objective of this study was to determine the molecular mechanism of HS effect on adipocyte differentiation and lipid storage in subcutaneous fat–derived porcine stromovascular cells (SVC).


MATERIALS AND METHODS

Primary Cell Isolation and Cell Culture

All animal care and protocols described in these experiments were approved by the Purdue Animal Care and Use Committee. Isolation and culture of primary porcine preadipocytes were based on the protocol described previously (Ramsay, 2005). Briefly, preadipocytes (SVC) were isolated from the inner layer of subcutaneous adipose tissue from male juvenile piglets (<7 d old). Excised tissue was immediately kept in buffered saline (0.15 M NaCl and 10 mM HEPES, pH 7.4) at 37°C. Tissue was thoroughly minced through repeated cutting with a pair of scissors and then incubated with a cocktail (10 mM NaHCO3, 10 mM HEPES, 5 mM d-glucose, 120 mM NaCl, 4.6 mM KCl, 1.25 mM CaCl2, 1.20 mM MgSO4, 1.20 mM KH2PO4, and 3% BSA) containing collagenase (Collagenase type I, 1 mg/mL; Worthington Biochemical Corp., Lakewood, NJ) in a shaking water bath at 37°C for 45 min at 120 oscillation/min. The digested sample was centrifuged at 2,000 × g for 10 min at 4°C to separate the floating adipocytes from the precipitated SVC pellet. The SVC pellet was washed twice in the digestion cocktail. Afterward, the pellet was suspended in Dulbecco’s modified Eagle medium/F12 (DMEM/F12) medium (Sigma-Aldrich, St. Louis, MO) with 10% fetal bovine serum (Mediatech, Manassas, VA) and 1% antibiotic–antimycotic (Sigma-Aldrich) and incubated in a humidified incubator with 5% CO2 and 95% air. After reaching confluence, cells were differentiated under normal (37°C) or HS (41.5°C) temperature environments for 3, 6, or 9 d in a differentiation medium: DMEM/F12 with 10% fetal bovine serum, 1% antibiotic–antimycotic, 1 μM insulin, 1 μM dexamethasone (1 mM), 1 μM rosiglitazone, 1 μM biotin, 1 μM triiodothyronine, and 1 μM pantothenic acid. The medium was replaced every 3 d and rosiglitazone was removed from the differentiation media after d 3 of differentiation. Cells showed visible signs of lipid accumulation from d 6 of differentiation and were fully lipid filled by d 9.

Oil Red O Staining

Oil red O staining was conducted following previously described protocol (Ramírez-Zacarias et al., 1992) with slight modifications. Briefly, cells in 24-well plates were washed twice with 1x PBS and immediately incubated in 10% formalin for 5 min at room temperature. The formalin was discarded and fresh formalin was added for another hour, after which cells were washed with 60% isopropanol and then incubated in oil red O working solution for 10 min. Cells were finally washed with deionized distilled water and photographed on a Nikon Eclipse TE 2000-U microscope (Nikon Instruments Inc., Melville, NY).

Gene Expression Analysis

Gene expression was measured through real-time PCR. Total RNA from adipocytes was extracted using QIAzol lysis reagent (Qiagen, Valencia, CA), based on the manufacturer’s protocol. Extracted RNA was dissolved in nuclease-free water (Ambion, Austin, TX). Concentrations of RNA were determined with a Nanodrop 1000 Spectrophotometer (Thermo Scientific, Waltham, MA). Integrity of RNA and genomic DNA contamination were checked by electrophoresis on 0.8% agarose. One microgram RNA was reverse transcribed with Moloney murine leukemia virus reverse transcriptase (Promega, Madison, WI). A Bio-Rad MyiQ thermocycler (Bio-Rad Laboratories, Inc., Temecula, CA) was used to conduct the PCR assays. The PCR reaction mixtures contained 0.5 μg of cDNA, 0.075 nmol of forward and reverse primers, respectively, and RT2 SYBR Green qPCR master mix (SABiosciences, Frederic, MD) in a total reaction volume of 20 μL. The reaction mix was incubated at 95°C for 5 min for the initial denaturation step and then 40 cycles of the following steps: 10 s at 95°C, 20 s at 55°C, and 20 s at 72°C. The sequence of real-time PCR primers is presented in Table 1. Abundance of mRNA for each gene was calculated after its cycle threshold (Ct) was normalized to the Ct for 18S using the ΔΔCt method.


View Full Table | Close Full ViewTable 1.

Primer sequences for real-time PCR

 
Gene1 Forward Reverse
CD36 5′–ATC GTG CCT ATC CTC TGG–3′ 5′–CCA GGC CAA GGA GGT TAA–3′
FATP4 5′–CAT TGT GGC TCA GCA GGT TA–3′ 5′–CAG GCT AGG GGT CAA ATC AA–3′
C/EBPα 5′–TGG ACA AGA ACA GCA ACG AG–3′ 5′– TTG TCA CTG GTC AGC TCC AG–3′
aP2 5′–TGG TAC AGG TGC AGA AGT GG–3′ 5′–ATT CTG GTA GCC GTG ACA CC–3′
HSP70 5′–TTC GTG GAC AGA AGC CAC AG –3′ 5′–TTG CTA GGA TCT CCA CCC GA–3′
FAS 5′–AGT TTG TGA TGG AGA ACA CGG CCT–3′ 5′–TGT TCA CAC GTG GTG CAA GGG TTA–3′
GK 5′–CGC TGA GGA AAG TGA AAT CCG–3′ 5′–TCG CGT CTT TGG AAT CTA CGA–3′
PCK1 5′–CCC TGC CTT TGA AAA AGC CC–3′ 5′–GGA GAT GAT TTC TCG GCG GT–3′
18S 5′–ATC CCT GAG AAG TTC CAG CA–3′ 5′–CCT CTT GGT GAG GTC GAT GT–3′
PPARγ 5′–GCC CTT CAC CAC TGT TGA TT–3′ 5′–GTT GGA AGG CTC TTC GTG AG–3′
LPL 5′–ATT CAC CAG AGG GTC ACC TG–3′ 5′–AGC CCT TTC TCA AAG GCT TC–3′
Leptin 5′–TTG GCC CTA TCT GTC CTA CG–3′ 5′–GTG ACC CTC TGT TTG GAG GA–3′
HSP27 5′–GAG CTG ACG GTC AAG ACC AA–3′ 5′–AAT GAA GCC GTG CTC ATC CT–3′
HSP90 5′–GTC GAA AAG GTG GTT GTG TCG–3′ 5′–TTT GCT GTC CAG CCG TAT GT–3′
GLUT4 5′–GAA GGA AGA AGG CAA TGC TG–3′ 5′–GAG GAA CCG TCC AAG AAT GA–3′
Adiponectin 5′–TTT CTG GGC CCA CTG TGT TT–3′ 5′–GGT TTT GCA TTG CAG GCT CA–3′
HSP60 5′–AGA TGC CCT GAA TGC GAC AA–3′ 5′–TGA CTC CAA GGC TGG AAT GC–3′
FATP6 5′–TTC TTC GGCTAT GCT GGC AA–3′ 5′–TGG ACC ATTAGG TCT CCG GT–3′
1aP2 = fatty acid binding protein 2; CD36 = fatty acid translocase 36; C/EBP α = CCAAT/enhancer binding proteins α; FATP4 = fatty acid transport protein 4; FATP6 = fatty acid transport protein 6; FAS = fatty acid synthase; GLUT4 = glucose transporter protein type 4; GK = glycerol kinase; HSP = heat shock protein; LPL = lipoprotein lipase; PCK1 = phosphoenolpyruvate carboxykinase 1; PPARγ = peroxisome proliferator-activated receptor γ.

Western Blot

Cells were initially washed twice with cold 1x PBS after which they were suspended in 1x radioimmunoprecipitation assay (RIPA) buffer (10% Nonidet P-40), 0.5 M Tris-HCl, 2.5% deoxycholic acid, 1.5 M HCl, and 10 mM EDTA) supplemented with commercial protease and phosphatase inhibitor cocktails (Sigma-Aldrich). Protein concentrations were determined by bicinchoninic acid assay (Thermo Fisher Scientific, St. Louis, MO). Equal amounts of protein were loaded on 10% SDS polyacrylamide gels. Proteins were transferred to nitrocellulose membranes (Bio-Rad Laboratories, Inc.). Thereafter, membranes were blotted with respective primary antibodies: anti-β-actin (Cell Signaling Technology, Danvers, MA), anti-heat shock protein (HSP) 70 (Cayman Chemicals, Ann Arbor, MI), anti-peroxisome proliferator-activated receptor γ (PPARγ) and anti-HSP90 (Cell Signaling Technology), anti-phosphoenolpyruvate carboxykinase 1 (PCK1 or PEPCK-C; Abcam Inc., Cambridge, MA), anti-glycerol kinase (GK; Bioss Inc., Woburn, MA), and anti-adiponectin antibody (Xeno Diagnostics, Indianapolis, IN).

Respective secondary antibodies were used (horseradish peroxidase [HRP]-conjugated goat anti-mouse or goat anti-rabbit IgG; Cell Signaling Technology). Membranes were developed with Immobilon HRP substrate (Millipore, Billerica, MA) and exposed to autoradiographic film (Santa Cruz Biotechnology, Dallas, TX). Band intensities were quantified with Kodak 1 D 3.6 software (Eastman Kodak Co., Rochester, NY).

Triglyceride Content Determination

Total cellular lipid was extracted with isopropanol for 30 min at room temperature. Thereafter, triglyceride (TAG) content was determined using a commercially available TAG determination kit (Sigma-Aldrich). Triglyceride content was normalized to cellular protein content.

Measurement of Cellular ATP Content

Concentration of ATP was measured in total cellular lysate recovered with 1x RIPA buffer. Levels of ATP were measured with an ATP determination kit (Invitrogen, Carlsbad, CA). Cellular ATP amount was normalized to cellular protein content.

Leptin ELISA

Cell culture medium was collected on d 6 of differentiation from adipocytes in both control and HS treatments. Leptin level was analyzed in duplicate wells with a pig leptin ELISA kit (CUSABIO, Wuhan, China) according to the manufacturer’s instruction. The minimum detectable limit of this assay was 0.25 ng/mL.

Statistical Analyses

Data were analyzed by ANOVA with GLM procedure and mean separation by Tukey analysis according to the procedures of SAS (SAS Inst. Inc., Cary, NC). Day and temperature were considered main effects and day × temperature was considered an interaction. Box–Cox transformation procedure was used when residuals were not normal. Differences were considered significant with P < 0.05, and P-values between 0.05 and 0.10 were considered as showing a strong tendency of significance. Values in the text are means ± SEM.


RESULTS

Regulation of Adipogenesis by Heat Stress

To test whether HS directly regulates adipocyte differentiation in the absence of other interfering in vivo factors, preadipocytes were subjected to differentiation over a 9-d period after which they were fully differentiated. Image from oil red O staining shows increased lipid staining of cells differentiated under HS compared with control cells (Fig. 1A). This is supported by the increased (P < 0.0001) content of TAG in cells differentiated in HS compared with control (approximately 3- and 1.6-fold greater) on d 6 and 9 of differentiation, respectively (Fig. 1B). Expression of key adipogenic transcription factors, PPARγ and CCAAT/enhancer binding proteins α (C/EBPα), were differentially regulated by HS such that HS did not affect the expression of PPARγ (P > 0.05) but led to an increase (P < 0.0001) in the expression of C/EBPα relative to the control (Fig. 2B). Immunoblotting of PPARγ protein also shows that treatment did not affect its protein abundance (Fig. 2C). Protein abundance of C/EBPα could not be determined due to the lack of a good antibody. It is unclear whether C/EBPα is regulated by the treatment at the protein expression level.

Figure 1.
Figure 1.

Heat stress (HS) stimulates adipogenesis in adipocytes. (A) oil red O staining of pig adipocytes on d 3, 6, and 9 of differentiation (40x) at 37 and 41.5°C temperatures. (B) Triglyceride (TAG) concentration was measured in pig adipocytes under normal conditions (37°C) and HS condition (41.5°C) on d 3, 6, and 9, respectively. Bars represent means ± SEM of at least 6 different replicates. *Mean difference between HS and control at P < 0.05.

 
Figure 2.
Figure 2.

Expression of genes related to adipogenesis in differentiating adipocytes under normal (37°C) and heat stress (41.5°C) conditions. (A) Peroxisome proliferator-activated receptor γ (PPARγ) gene expression. (B) CCAAT/enhancer binding proteins α (C/EBPα) gene expression. (C) PPARγ immunoblotting (upper panel) and quantification (lower panel). Bars represent means ± SEM of at least 6 different replicates. *Mean difference between heat stress and control at P < 0.05.

 

Heat Stress Promotes Increased Expression of Fatty Acid Transport and Glyceroneogenic Genes

Expression of several other genes implicated in increased TAG storage such as fatty acid synthase (FAS), LPL, adipocyte protein 2 (FABP4 or aP2), fatty acid translocase 36 (CD36), fatty acid transport protein 6 (FATP6) and glucose transporter protein type 4 (GLUT4) were significantly induced by HS compared with control (Fig. 3). Unlike FATP6, expression of fatty acid transport protein 4 (FATP4) was not different between control and HS treatments (Fig. 3E and F). Gene expression of PCK1 and GK was also measured. These are key enzymes involved in glyceroneogenesis, a pathway for making the glycerol backbone of TAG. Heat stress increased (P < 0.0001) the transcript levels of both PCK1 and GK (Fig. 4A and 4B) and their protein abundance (Fig. 4C and 4D).

Figure 3.
Figure 3.

Expression of genes involved in fatty acid transport and binding and glucose transport in adipocytes under normal condition (37°C) and heat stress condition (41.5°C) on d 3, 6, and 9, respectively. Genes were measured by real-time PCR. (A) Fatty acid synthase (FAS). (B) Adipocyte protein 2 (aP2). (C) Fatty acid translocase 36 (CD36). (D) Lipoprotein lipase (LPL). (E) Fatty acid transport protein 4 (FATP4). (F) Fatty acid transport protein 6 (FATP6). (G) Glucose transporter protein type 4 (GLUT4). Bars represent means ± SEM of at least 6 different replicates. *Mean difference between heat stress and control at P < 0.05.

 
Figure 4.
Figure 4.

Heat stress increases abundance of genes involved in glyceroneogenesis in adipocytes. (A) Phosphoenolpyruvate carboxykinase 1 (PCK1 or PEPCK-C) and (B) glycerol kinase (GK) mRNA levels were measured by real-time PCR. (C) and (D) are immunoblots for PCK1 and GK, respectively. Bars represent means ± SEM of at least 6 different replicates. *Mean difference between heat stress and control at P < 0.05.

 

Increased Expression of Adipokines in Heat Stress

Leptin and adiponectin play major paracrine, endocrine, and autocrine roles in the regulation of metabolism (Lee and Shao, 2014; Blüher and Mantzoros, 2015) and may be implicated in the alteration of metabolism observed in pigs under HS. To test this hypothesis, we measured both leptin and adiponectin expression. Expression of both adipokines was increased (P < 0.05) in HS (Fig. 5A through 5E), suggesting that they may play critical roles during HS.

Figure 5.
Figure 5.

Heat stress stimulates adipokines expression in pig adipocytes. (A) Leptin and (B) adiponectin mRNA abundance was measured by real-time PCR in adipocytes under normal (37°C) and heat stress (41.5°C) conditions. (C) Media leptin was measured by ELISA. (D) Media adiponectin was measured by western blotting with quantification. Bars represent means ± SEM of at least 6 different replicates. *Mean difference between heat stress and control at P < 0.05.

 

Heat Stress Promotes Expression of Heat Shock Proteins in Pig Adipocytes

Heat shock proteins, molecular chaperones against the damaging effects of HS, were also quantified. The transcript levels of multiple HSP (HSP27, HSP60, HSP70, and HSP90) were greater (P < 0.0001) in HS compared with control (Fig. 6). Western blot analysis of HSP70, the most important cytoplasmic HSP, and HSP90 also indicated that HS increased their protein abundance (Fig. 6E and 6F). Therefore, the induction of HSP plays a role in the adaptive response against mild HS in pig adipocytes.

Figure 6.
Figure 6.

Heat stress stimulates heat shock protein (HSP) expressions in pig adipocytes. Transcript levels of HSP 27(A), HSP60 (B), HSP70 (C) and HSP90 (D) were measured in pig adipocytes under normal (37°C) and heat stress (41.5°C) conditions using real-times PCR. (E) HSP70 and (F) HSP90 protein expression was determined by western blot. Bars represent means ± SEM of at least 6 different replicates. *Mean difference between heat stress and control at P < 0.05.

 

Heat Stress Leads to Increase in Cellular ATP Level

Mild HS is able to activate fight-or-flight reactions, which need the cellular ATP pool as an energy source. Therefore, we measured cellular ATP levels in the differentiating adipocytes. As expected, total cellular ATP decreased with the progression of differentiation (Fig. 7). However, cells in HS had greater (P < 0.001) cellular ATP than those in control temperature.

Figure 7.
Figure 7.

Heat stress leads to increase cellular ATP concentration. Adenosine triphosphate level was measured in pig adipocytes under normal (37°C) and heat stress (41.5°C) conditions. Bars represent means ± SEM of at least 6 different replicates. *Mean difference between heat stress and control at P < 0.05.

 


DISCUSSION

Pigs have been known to conserve adipose tissue mass during HS (Collin et al., 2001; Kouba et al., 2001), resulting in increased adiposity, especially during the hot summer months. Previous studies have implicated the upregulation of LPL (Kouba et al., 2001) and FAS activities (Pearce et al., 2013a) in adipose tissue of pigs under mild HS condition. Increased LPL activity promotes greater hydrolysis of TAG in serum lipoproteins into fatty acids and glycerol, and these are then reesterified into TAG in adipocytes, resulting in increased adipose tissue mass. Both LPL and FAS are transcriptionally regulated (Wolf et al., 1994; Zechner et al., 2000; Wang and Eckel, 2009) by insulin action, and this suggests that the increased expressions of these genes in HS may be tied to the elevated insulin concentrations observed in pigs under HS (Pearce et al., 2013a). However, a dominant role for elevated insulin is questionable (Williams et al., 2013; Pearce et al., 2014), given that HS in pigs is not always associated with elevated circulating serum insulin concentration, suggesting that additional mechanisms may be involved. In this study, we have applied an in vitro model of adipocyte differentiation to determine the direct effects of HS on porcine adipose tissue metabolism. This model provides a tool for direct investigation of effects of the elevated temperatures without the complications of in vivo factors such as variable insulin, glucocorticoids or catecholamine concentrations, or the potential involvement of other tissues.

Results from the current study indicate that HS stimulates adipogenesis in pig adipocytes and it is accompanied by increased TAG storage. Multiple genes involved in increased adipogenesis are induced by HS. Increased expression of adipogenic transcription factor, C/EBPα, suggests that it may be implicated in the increased adipogenesis under HS. However, the mRNA and protein abundance of PPARγ were not affected by HS. This indicates that HS may not directly regulate its expression. In addition, genes that are involved in increased fatty acid uptake (aP2, CD36, LPL, and FATP6), de novo fatty acid synthesis (FAS), glucose uptake (GLUT4), and glyceroneogenesis (PCK1 and GK) were induced. The induction of these genes in HS may indeed reflect an increased cooperativity between C/EBPα and PPARγ during HS, because most of these genes are under transcriptional regulation by C/EBPα and PPARγ (Lin and Lane, 1994; MacDougald and Lane, 1995). Increased expression of the fatty acid transporters, CD36 and FATP6, agrees with the expected increas in in free fatty acids availability, presumably from an increased LPL activity. To support this idea, there is evidence that FATP6 and CD36 colocalize to enhance long-chain fatty acid uptake, an interaction that is not seen between CD36 and FATP4 (Gimeno et al., 2003). Increased expression of these fatty acid transporting genes may partly explain the reduction in serum fatty acid concentration in pigs under HS (Pearce et al., 2013a).

Induction of glyceroneogenic genes, PCK1 and GK, also provides another mechanistic explanation for the increased TAG storage in adipose tissue during HS. With the increased influx of free fatty acids into adipocytes, increased esterification of these acids is warranted to prevent lipotoxicity (Kawai et al., 2001). Both PCK1 and GK are directly involved in glyceroneogenesis in adipocytes, leading to the production of glycerol backbone that is critical for fatty acid condensation via glycerol-3-phosphate (Reshef et al., 2003). It is quite interesting that, compared with unstimulated mouse white adipocytes (Guan et al., 2002; MacDougald and Burant, 2005), pig adipocytes express significant amount of GK and its expression is inducible by HS. Therefore, both PCK1 and GK may play important roles in the increased lipid storage in pig adipocytes during HS.

Increased expression of adipokines (leptin and adiponectin) in adipocytes under HS is consistent with the report of others that HS stimulates secretion and expression of leptin and adiponectin in 3T3-L1 adipocyte (Bernabucci et al., 2009) and mice (Morera et al., 2012). Leptin, an adipokine that is expressed exclusively in adipose tissue, is a lipostat that plays a major role in appetite control through a feedback effect on key areas of the hypothalamus, ultimately leading to suppression of feed intake (Minokoshi et al., 2002). As we have shown previously, leptin administration results in reduction of feed intake in pigs (Ajuwon et al., 2003). A major adaptation to HS in pigs is the reduction in feed intake (Pearce et al., 2013a). Therefore, it is conceivable that elevated leptin concentration from adipocytes in pigs under HS may play an anorexigenic role in the animals leading to suppression of feed intake (Minokoshi et al., 2002).

Increased expression of HSP (HSP27, HSP60, HSP70, and HSP90) supports the hypothesis that nonlethal HS will result in appropriate adaptive response directly in pig adipocytes, comparable to increased tissue expression of HSP in pigs under HS (Pearce et al., 2013b). Although HSP are known to be involved in restoring cellular functions and enhancing cellular survivability during heat and other stresses (Horowitz and Robinson, 2007), their role in the increased lipid storage in adipocytes under HS is unknown. Heat shock protein 27 is an actin polymerization factor and its phosphorylation status promotes the polymerization of actin under HS conditions (Lavoie et al., 1995; Guay et al., 1997). Mechanistically, PPARγ could activate HSP22, which is responsible for activating HSP27 (Sun et al., 2004; Hamza et al., 2009). Furthermore, it is known that HSP90 acts as a chaperone for PPARγ, leading to increase its stability under stress (Nguyen et al., 2013), and there is evidence that HSP90 may be involved in the enhancement of the phosphoinositide-3-kinase–protein kinase B/Akt (PI3K-PKB/Akt) pathway under HS (Sato et al., 2000), a pathway that is involved in the stimulation of glucose uptake and GLUT4 translocation in adipocyte (Kohn et al., 1996). It is also known that HS stimulates HSP70 accumulation on adipocyte lipid droplets to stabilize the droplet monolayer or chaperone denatured proteins for subsequent refolding (Jiang et al., 2007).

Whether HSP are involved in other functions to increase TAG storage in adipocytes is unknown. Nevertheless, based on their established function, induction of HSP in the adipocytes would be expected to serve a protective role, preventing the side effects of HS, such as ensuring appropriate protein folding, preventing protein aggregation, and refolding them into functional native states.

Heat stress response is an energy demanding process, with greater ATP requirement for increasing expression and activity of HSP. The reduction in cellular ATP with the progression of adipocyte differentiation in this study is consistent with earlier reports showing reduced cellular ATP concentration with adipocyte differentiation (Luo et al., 2008). However, HS appears to preserve cellular ATP compared with cells at 37°C. This agrees with the findings of Jayakumar et al. (1999) in myocytes that HS could help preserve cellular ATP levels through alteration in energy metabolism profile, perhaps leading to increased production of ATP to support the HS response.

In conclusion, we provide evidence herein that HS directly leads to increased adipogenesis in porcine adipocytes. The mechanism of this response may include increased expression of transcription factor C/EBPα and increased expression of fatty acid transport and glyceroneogenic genes. Increased expression of leptin in adipocytes under HS may partly explain the reduction in feed intake observed in pigs under HS. These direct observations in adipocytes offer useful insights into the metabolic response in pig adipose tissue during HS and provide plausible mechanisms underlying the increased adipose tissue mass long observed in pigs under HS.

 

References

Footnotes


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