J. Anim. Sci. 2005. 83:565-578
© 2005 American Society of Animal Science
ANIMAL GROWTH, PHYSIOLOGY, AND REPRODUCTION |
Expression of adiponectin and its receptors in swine1,2
E. Lord*,
S. Ledoux*,
B. D. Murphy*,
D. Beaudry
and
M. F. Palin,3
* Centre de Recherche en Reproduction Animale, Faculté de Médecine Vétérinaire, Université de Montréal, St-Hyacinthe, Québec, Canada J2S 7C6; and
and
Dairy and Swine Research and Development Center, Agriculture and Agri-Food Canada, Lennoxville, Québec, Canada J1M 1Z3
 |
Abstract
|
|---|
Adiponectin is an adipocyte-derived hormone that plays an important role in lipid metabolism and glucose homeostasis. Objectives of this study were 1) to determine the presence and distribution of adiponectin and its receptors 1 and 2 (adipoR1 and adipoR2) in porcine tissues; 2) to characterize pig adiponectin, adipoR1, and adipoR2 mRNA levels in various fat depots from three different breeds of pigs; and 3) to study, in stromal-vascular cell culture, the effects of leptin and tumor necrosis factor-
(TNF) on pig adiponectin, adipoR1, and adipoR2 gene expression. To this end, fat Chinese Upton Meishan (UM, n = 10), lean Ham Line (HL, n = 10), and Large White (LW, n = 10) gilts were used. We report the isolation of partial cDNA sequences of pig adipoR1 and adipoR2. Porcine-deduced AA sequences share 97 to 100% homology with human and murine sequences. Pig adipoR1 mRNA is abundant in skeletal muscle, visceral fat, and s.c. fat tissues, whereas adipoR2 mRNA is predominantly expressed in liver, heart, skeletal muscle, and visceral and s.c. fat tissues. Pig adiponectin mRNA levels in s.c. and visceral fat tissues were not associated with plasma insulin and glucose in fasting animals. Subcutaneous (r = –0.44, P < 0.05), visceral (r = –0.43, P < 0.05), and total body fat (r = –0.42, P < 0.05) weights were negatively correlated with adiponectin mRNA levels measured in visceral, but not s.c., fat. Pig adipoR1 and adipoR2 mRNA levels, in visceral fat, were less expressed in fat UM gilts than in the lean HL gilts (P < 0.05). Inverse associations were found between s.c. (r = –0.57, P < 0.01), visceral (r = –0.46, P < 0.05), and total body fat (r = –0.56, P < 0.01) weights and adipoR2 mRNA levels in visceral fat only. We were unable to find such associations for adipoR1 mRNA levels in the overall gilt population. The current study demonstrated that TNF downregulates adiponectin and adipoR2, but not adi-poR1, mRNA levels in stromal-vascular cell culture. Moreover, leptin significantly decreased adiponectin mRNA levels, whereas there was no effect on adiponectin receptors. We conclude that adiponectin and adi-poR2 mRNA levels, but not adipoR1, are modulated in pig visceral fat tissues. Furthermore, our results indicate that TNF interferes with adiponectin function by downregulation of adipoR2 but not of adipoR1 mRNA levels in pigs.
Key Words: Adiponectin • Adiponectin Receptors • Fat • mRNA Levels • Pig
|
Introduction
|
|---|
Adipose tissue, well known for its function as an energy reservoir, is now recognized as an endocrine organ that produces numerous proteins with broad biological activities. These include IL-6, tumor necrosis factor- (TNF), leptin (Coppack, 2001
), resistin (Steppan et al., 2001), and adiponectin (Maeda et al., 1996). Circulating adiponectin levels are decreased in obese human subjects (Arita et al., 1999) and in individuals with type 2 diabetes and coronary artery disease (Hotta et al., 2000). Administration of adiponectin lowers circulating glucose and ameliorates insulin resistance in mice (Yamauchi et al., 2001), and adiponectin-deficient mice develop insulin resistance and diabetes (Kubota et al., 2002; Maeda et al., 2002). In response to the binding of adiponectin, adiponectin receptors 1 (adipoR1) and 2 (adipoR2) mediate intracellular signalling events, including increased adenosine 5'-monophosphate kinase and peroxisome proliferator-activator receptor- ligand activity, with the consequence of increased fatty acid oxidation and glucose uptake (Yamauchi et al. 2003). Adiponectin expression can be modulated by TNF, a pro-inflammatory cytokine (Fasshauer et al., 2002). Higher levels of TNF mRNA and protein are associated with obesity in humans and pigs (Hotamisligil et al., 1999; Chen et al., 2004). Moreover, TNF was the first adipocytokine proposed to represent a molecular link between obesity and insulin resistance (Moller, 2000). The purposes of this study were to establish the existence of adiponectin receptors in the pigs, to determine tissue distribution of adiponectin and its receptors, and to establish whether the relative expression of adiponectin and its receptors, in s.c. and abdominal fat depots, varies among divergent porcine genotypes. Finally, the effects of TNF and leptin on the expression levels of adiponectin, adipoR1, and adipoR2 mRNA were further studied during the differentiation process of porcine stromal-vascular (SV) cells to adipocytes.
|
Materials and Methods
|
|---|
Animals and Tissue Collection
A total of 30 gilts, kindly provided by Hypor (Regina, Saskatchewan, Canada), were used in this study. These included Chinese Upton Meishan (UM, n = 10), Large White (LW, n = 10) and Ham Line (HL, n = 10) gilts. These breeds differ considerably in body composition and body fat distribution (shown in Table 1). Piglets were transported to the research center on reaching 15 ± 2 kg of BW. They were fed a commercial diet (50% corn, 20% barley, 20% wheat bran, and 5% soybean; as-fed basis) ad libitum until slaughter at 109.9 ± 2.96 kg of live weight. Animals had free access to fresh water during the entire trial. A crossbred adult sow (Duroc x Yorkshire x Landrace) was sampled to establish adiponectin and adiponectin receptor tissue distribution.
At slaughter, live weight was taken, and backfat and abdominal visceral fat tissue samples were collected and immediately frozen in liquid nitrogen. Jugular blood samples were collected from gilts to measure fasting insulin and glucose. Various tissue samples, including s.c. fat, visceral fat, lung, kidney, stomach, skeletal muscle, liver, ovary, uterus, brain and heart, were collected for tissue distribution studies. All tissue samples were frozen immediately in liquid N and stored at –80°C until needed. After evisceration, abdominal visceral fat was completely removed from the carcasses and weighed. Total s.c. fat weight (kg) was measured by carcass dissection according to Marcoux et al. (2003), with the exception that the s.c. fat weight included the skin. One side of each carcass was separated into four cuts (shoulder, ham, belly, and loin), and each part was completely dissected to measure the complete carcass s.c. fat. Total body fat was obtained by adding the abdominal visceral fat to the s.c. fat and intermuscular fat (kg). Animals were cared for according to the recommended code of practice and killed using an acceptable method approved by the local Animal Care Committee following the guideline of the CCAC (1993).
Biochemical Analyses
Plasma glucose was measured by colorimetry using a commercial kit (No. 1448668; Roche Diagnostic Corp., Indianapolis, IN) and according to the manufacturer’s instructions. Glucose intra- and interassay CV were 0.7 and 1.0%, respectively. Plasma insulin was measured by RIA (catalog No. 11002, Immunocorp, Montreal, Québec, Canada). The intra- and interassay CV for the insulin RIA were 4.1 and 4.7%, respectively. Assay sensitivity was 26.9 pmol/L.
Stromal-Vascular Cell Culture
Dorsal s.c. adipose tissue was aseptically removed from five crossbreed piglets at 5 d of age, and SV cells were isolated according to a method used by Ramsay (2001). Briefly, adipose tissue was minced with scissors and incubated for 90 min in a digestion buffer made of Dulbecco’s modified Eagle’s medium (DMEM)/F12 (Sigma, Oakville, Ontario, Canada), 100 mM HEPES, and 1.5% BSA at pH 7.4, which contained 2 mg/mL of Type I collagenase/g of tissue (Gibco BRL, Burlington, Ontario, Canada). Incubation was performed at 37°C in a shaking agitator. A fivefold excess of digestion buffer, without collagenase, was then added to the digested adipose tissue solution and the resulting solution was then filtered through 250- and 20-µm nylon mesh (Sefar America, Depew, NY). Cells where then centrifuged at 1,000 x g for 10 min at 4°C. Supernatant fluid was discarded and cells were resuspended in plating medium made of DMEM/F12, 10% fetal bovine serum (Gibco-Invitrogen, Grand Island, NY), 80 nM dexamethasone (Sigma), and antibiotics: 40 mg/L of gentamicin sulfate (Sigma), 50 mg/L of cephalothin (Sigma), and 2 mg/L of fungizone (BioWhittaker, Walkersville, MD). Aliquots of SV cells were stained with 0.4% trypan blue and counted on a hemocytometer to evaluate the number of viable/dead cells. Cells were plated at a density of 5 x 104 cells/cm2 in 24-well culture plates. They were maintained at 37°C in a humidified, 5% CO2 atmosphere until they reached confluence. At that time, medium was changed to serum-free medium for 24 h. After this starvation period, cultures were then changed to a serum-free media that contained DMEM/F12 and antibiotics. These cell cultures were then treated with 1) a combination of 850 nM insulin, 2.75 µg/mL of transferin, and 2.5 µg/mL of selenium (ITS; Sigma); 2) ITS + 10 ng/mL of recombinant human TNF (Sigma); or 3) ITS + 100 ng/mL of recombinant porcine leptin (Ruiz-Cortes et al., 2003). We used ITS to induce differentiation of preadipocytes to adipocytes because it was previously demonstrated that insulin is sufficient to promote differentiation of porcine SV cells to adipocytes (Hausman, 1989; Boone et al., 2000). Culture medium was changed every 2 d. Cells from each treatment group were collected at 0, 24, 48, 72, and 96 h (two wells collected at each time) and immediately frozen at –80°C for RNA isolation. Stromal-vascular cell cultures were stained for lipids using Oil Red O as described by Magun et al. (1996) and were counterstained with Giemsa to evaluate cellular proliferation and differentiation. The extent of differentiation was estimated by measuring adipocyte-specific fatty acid-binding protein (aP2) mRNA levels by reverse-transcriptase PCR (RT-PCR) analysis as previously reported (Ding et al. 1999). Oligonucleotides and PCR conditions used to amplify porcine aP2 are described in Table 2.
View this table:
[in this window]
[in a new window]
|
Table 2. Primer sequences and optimal conditions for PCR of porcine adiponectin, adiponectin receptor 1 and 2 (adipoR1, adipoR2), cyclophilin, and adipocyte-specific fatty acid-binding protein (aP2) genes
|
|
RNA Extraction and cDNA Preparation
Total RNA was extracted from collected tissue samples using TRIzol reagent (Gibco BRL) according to the manufacturer’s instructions. The RNA isolation for SV cell culture was done using RNeasy mini kit (Qiagen, Mississauga, Ontario, Canada) according to the manufacturer instructions. The extracted RNA was dissolved in water and quantified spectrophotometrically at 260 nm. For each sample, an RNA aliquot was subjected to electrophoresis in a 1% agarose gel to verify its integrity. Total RNA was reverse transcribed to cDNA in a PTC-200 Peltier programmable thermal cycler (MJ Research, Foster City, CA). Five micrograms of total RNA was treated with 3 U of DNase I (Gibco BRL) to remove contaminating genomic DNA. For the abdominal visceral fat samples, 2 µg of total RNA was treated with 2 U of DNase I. For the SV cells, 1 µg of total RNA was treated with 1 U of DNase I. The first strand cDNA was synthesized using the SuperScriptTM II preamplification system (Gibco BRL) and 500 ng of oligo(dT)12-18 as a primer (Amersham Pharmacia Biotech, Baie D’Urfé, Québec, Canada) in 50 µL of total reaction volume.
Cloning and Sequencing of Porcine Adiponectin Receptors
To determine porcine specific sequences of adipoR1 and adipoR2, primers were designed for PCR amplifications based on homology between human (Accession No. NM_015999 for R1; NM_024551/BC051858 for R2) and murine (Accession No. BC014875 for R1) sequences. For both receptors, 2.5 µg of total RNA from porcine s.c. adipose tissue was reverse-transcribed using Superscript RNase H reverse transcriptase (Life Technologies) according to the manufacturer’s instructions. For adipoR1, the PCR reaction was performed using primers R1-A and R1-1 (Table 2
) and the appropriate PCR conditions. A 1,044-bp product was gel-extracted using the QIAquick gel extraction kit (Qiagen) and according to the manufacturer’s instructions. Porcine adipoR1 was sequenced (three independent PCR amplifications) using the Big Dye terminator cycle sequencing ready reactions (PE Applied Biosystems, Foster City, CA) according to manufacturer’s instructions, and run on an ABI 377 (PE Applied Biosystems). A partial porcine adipoR1 coding a 1,044-bp sequence was submitted to GenBank (Accession No. AY452710). For adipoR2, the PCR reaction was performed using primers R2-A and R2-1 and the PCR conditions indicated in Table 2. A 652-bp fragment was gel-extracted using the QIAquick gel extraction kit (Qiagen) and according to the manufacturer’s instructions. Porcine adipoR2 was sequenced as described above for porcine adipoR1 (three independent PCR amplifications). The partial porcine adipoR2 coding sequence of 652 bp was submitted to GenBank (Accession No. AY452711).
Tissue Distribution of Porcine Adiponectin and Adiponectin Receptors 1 and 2
The PCR amplifications of adiponectin and adipoR1 and adipoR2 cDNA were performed in various pig tissues. The PCR reactions were performed using primers Adipo1 and Adipo2 for adiponectin, primers Adipor1-1 and Adipor1-2 for adipoR1 and primers Adipor2-1 and Adipor2-2 for adipoR2 (Table 2). These PCR reactions were performed in a 100-µL total volume, which contained 2 µL of the reverse transcriptase product, 200 µM deoxyribonucleotide triphosphate, 0.5 U of Taq polymerase in 1x Taq polymerase buffer (Amersham Pharmacia Biotech), and their corresponding concentrations of MgCl2 and forward and reverse primers (Table 2). The PCR profiles consisted of an initial denaturation step at 94°C for 2 min, followed by a fixed cycle number (Table 2) of denaturing at 94°C for 1 min, followed by the corresponding annealing temperature for 1 min (Table 2), extension at 72°C for 1 min and a final extension at 72°C for 5 min.
Pig cyclophilin amplification was also performed, using primers Cyclo1 and Cyclo2 (Table 2), to have an internal control for variations in cDNA synthesis. The 100-µL PCR reaction mixture contained 2 µL of the reverse transcriptase product, 200 µM deoxyribonucleotide triphosphate, 0.5 U of Taq polymerase in 1x Taq polymerase buffer (Amersham Pharmacia Biotech), and optimal concentrations of MgCl2 and forward and reverse primers (Table 2). The PCR profile used for cyclophilin was the same as described above for adiponectin. Adiponectin, adipoR1, adipoR2, and cyclophilin-amplified PCR fragments were separated on a 1% agarose gel and stained with ethidium bromide. Pictures of resulting gels were taken with Polaroid (Mississauga, Onatrio, Canada) positive-negative film. Negatives were then scanned using an imaging densitometer (model GS-670, Bio-Rad Laboratories Ltd., Mississauga, Ontario, Canada). The PCR amplifications were repeated in duplicates using two different tissue samples from the same animal.
Quantitative Measurements of Adiponectin and Adiponectin Receptor mRNA Expression in Fat Tissues
Subcutaneous fat and abdominal visceral fat cDNA were analyzed for adiponectin, adipoR1, and adipoR2 mRNA levels using real-time PCR amplification. For adiponectin, the forward 5'-ATGATGTCACCACTGGCAAATTC-3' and reverse 5'-GACCGTGACGTGGAAGGAGA-3' primers were designed based on the sequence of porcine adiponectin mRNA (GenBank Accession No. AY135647) and were selected with Primer Express Software (PE Applied BioSystem). Real-time PCR amplifications were performed in a 25-µL reaction volume consisting of 900 nM forward primer, 300 nM reverse primer, 1 µL of cDNA, 0.25 µL of AmpErase (PE Applied Biosystems), and 1x SYBR Green Master Mix (PE Applied Biosystems). Cycling conditions were 2 min at 50°C, followed by 10 min at 95°C. Then, 40 repetitive cycles of 15 s at 95°C and 1 min at 60°C, were performed. All procedures were performed on an ABI Prism 7700 sequence detector (PE Applied Biosystems). The adipoR1 was amplified in real time using forward 5'-GCCATGGAGAAGATGGAGGA-3' and reverse 5'-AG-CACGTCGTACGGGATGA-3' primers, designed from the sequence of porcine adipoR1 mRNA (GenBank Accession No. AY452710). Real-time PCR amplifications of adipoR1 were performed using the same conditions as previously described for adiponectin except that 300 nM and 50 nM of forward and reverse primers were used, respectively. AdipoR2 real-time amplifications were performed using forward 5'-TGTTCGCCACCCC-TCAGTAT-3' and reverse 5'-AATGATTCCACTCAGGCCCA-3' primers, based on adipoR2 porcine mRNA sequence (GenBank Accession No. AY452711). The adipoR2 real-time PCR amplification conditions were identical to those described above for adiponectin, with the exception of forward and reverse primers quantities, which was 50 nM for both primers. Samples were normalized using the housekeeping gene cyclophilin (primer sequences; 5'-GCA CTG GTG GCA AGT CCA T-3' and 5'-AGG ACC CGT ATG CTT CAG GA-3', Gen-Bank Accession No. AY266299). Cyclophilin real-time PCR amplifications were performed using the same conditions except that 300 nM of each forward and reverse primer were used. The PCR amplifications were performed in triplicate. Standard curves were prepared in duplicate, for adiponectin, adipoR1, adipoR2, and cyclophilin. A pool of visceral and s.c. fat tissues cDNA was used to create a standard curve for quantification of the transcripts using the relative standard curve method as described by Applied Biosystems (1997). Standard curve arbitrary units were set at 1 for the undiluted cDNA pool, and n-fold dilutions of 0.75, 0.50, 0.25, 0.10, 0.05, 0.025, and 0.005 were then performed. The relative mRNA levels of each studied genes and endogenous cyclophilin reference were determined by interpolating the threshold cycle values to their respective standard curves. Specificity of the amplified fragments was verified on 3% agarose gel and with the melting curve analysis.
Quantitative Measurements of Adiponectin and Adiponectin Receptors mRNA Expression in Stromal-Vascular Cell Culture
Stromal-vascular cell culture cDNA were analyzed for adiponectin, adipoR1, and adipoR2 mRNA levels. Primers were those described above. Real-time PCR amplifications were performed in a 25-µL reaction volume consisting of 50 nM forward primer, 50 nM reverse primer, 1 µL of cDNA, 0.25 µL of AmpErase (PE Applied Biosystems), and 1x SYBR Green Master Mix (PE Applied Biosystems) for each of these genes. Samples were normalized using cyclophilin housekeeping gene. Cyclophilin RT-PCR amplifications were performed using the same conditions, except that 50 and 300 nM forward and reverse primers were used, respectively. A pool of SV cDNA was used to create a standard curve for quantification of the transcripts using the relative standard curve method as described above. The PCR amplifications were performed in triplicate.
Statistical Analyses
Relative quantification of mRNA levels was performed according to the standard curve method described by Applied Biosystems (1997). Differences among breeds were analyzed using one-way ANOVA followed by Dunnett adjustment for comparison to the UM reference group. Data were presented as mRNA quantities relative to the UM reference group with their SEM. Differences between treatments in SV cell culture were analyzed by one-way ANOVA with repeated measures in time. Correlation analysis between adiponectin, adipoR1, and adipoR2 mRNA levels and carcasses traits was calculated using the PROC CORR procedure of SAS (Version 8.1, SAS Inst., Inc., Cary, NC). Statistical significance was set at P < 0.05.
|
Results
|
|---|
Sequencing of Adiponectin Receptors 1 and 2 Reverse-Transcriptase PCR Products
Using RT-PCR on total RNA isolated from porcine adipose tissue, partial cDNA coding sequences were obtained for adipoR1 and adipoR2 (GenBank Accession No. AY452710 and AY452711, respectively). Analysis of the porcine adipoR1 nucleotide sequence showed that it is 92% identical to both the human and murine sequences (Yamauchi et al., 2003). Similarity analysis of adipoR1-deduced AA sequence showed 98 and 97% identity when compared with the human and mouse sequences, respectively (Figure 1A). The porcine adipoR2 displayed a nucleotide sequence that was 92 and 90% identical to the human and murine sequences, respectively (Yamauchi et al., 2003). Analysis of the deduced adipoR2 AA sequence showed a 100 and 99.5% identity compared with these same species (Figure 1B). Hydropathy plots of the porcine adipoR1 and adipoR2 sequences indicate that both have seven transmembrane domains along with a cytoplasmic amino-terminal domain and extracellular carboxyl-terminal (Figure 1). The adipoR1 and adipoR2 are highly related proteins in the mouse, and porcine adipoR1 and adipoR2 available sequences also are highly related and share 77% identity.
View larger version (81K):
[in this window]
[in a new window]
|
Figure 1. Deduced AA sequence of pig adiponectin receptors 1 and 2. A) Alignment of the predicted AA sequences of pig (AY452710), human (NM_015999), and mouse (BC014875) adiponectin receptor 1 (adipoR1). B) Alignment of the predicted AA sequences of pig (AY452711), human (BC051858), and mouse (AY424291) adiponectin receptor 2 (adipoR2). The hypervariable region of adiponectin receptor 1 is underlined and the phosphorylatable threonine is in bold character. Numbers indicate AA residue position. Hyphens are gaps introduced to optimize alignment. The seven transmembrane domains are shaded.
|
|
Effect of Breed on Plasma Glucose and Insulin, Body Weight, and Carcass Fat Distribution
At slaughter, total body fat, visceral fat, and s.c. fat were higher in UM gilts than in HL and LW gilts (P < 0.001; Table 1). There were no breed effects associated with fasting plasma insulin and glucose concentrations (Table 1).
Tissue Distribution of Pig Adiponectin and Adiponectin Receptors 1 and 2 mRNA
Expression of adiponectin, adipoR1, and adipoR2 mRNA was assessed by RT-PCR analyses in various tissues from an adult sow. The expected 709-bp fragment was amplified for pig adiponectin in adipose tissues (Figure 2), but it was not found in liver, kidney, heart, lung, ovary, stomach, brain, or uterus. A faint band corresponding to the expected length was also detected in skeletal muscle, presumably owing to the presence of i.m. fat in that tissue. A single 344-bp band for adipoR1 was amplified in all pig tissues tested, with the most abundant mRNA expression occurring in skeletal and adipose tissues (Figure 2). The 650-bp adipoR2 fragment was also expressed ubiquitously in the pig, with the greatest abundance found in the liver, heart, skeletal muscle, and adipose (Figure 2).
View larger version (42K):
[in this window]
[in a new window]
|
Figure 2. Tissue distribution of adiponectin and adiponectin receptors 1 and 2 mRNA in various tissues collected from an adult sow. The tissues were as follows: Lane 1 = liver; Line 2 = kidney; Lane 3 = heart; Lane 4 = lung; Lane 5 = ovary; Lane 6 = stomach; Lane 7 = brain; Lane 8 = uterus; Lane 9 = skeletal muscle; Lane 10 = ham s.c. fat; Lane 11 = back s.c. fat; Lane 12 = visceral fat; (–) = PCR amplification negative control without cDNA; Kb+ = Kb plus DNA Ladder (Invitrogen, Grand Island, NY). Expected fragment length (bp) is indicated on the right. Equal amounts of PCR products were loaded per lane as adjusted by cyclophilin housekeeping gene.
|
|
Expression of Pig Adiponectin mRNA in Subcutaneous and Visceral Fat Tissues
Using real-time PCR, pig adiponectin mRNA levels were quantified in s.c. and visceral fat tissues from different breeds of pigs. In the s.c. tissue, levels of adiponectin mRNA were higher in HL than in UM gilts (P < 0.05; Figure 3A). In visceral fat tissue, no significant differences were observed in adiponectin mRNA levels for these breeds (Figure 3B). Real-time PCR analysis also revealed that the expression of adiponectin mRNA was 52, 69, and 88% higher in visceral than s.c. fat tissue in the HL, LW, and UM breeds, respectively (data not shown).
View larger version (14K):
[in this window]
[in a new window]
|
Figure 3. Relative abundance of adiponectin mRNA in s.c. (A) and visceral (B) adipose tissues. Values (±SEM) are expressed as mRNA quantities relative to the UM reference group, which was given a value of one. Ham Line = HL (n = 10), Large White = LW, (n = 10), and Upton-Meishan = UM (n = 10). The asterisk indicates that means differed, P < 0.05.
|
|
Correlation analyses between adiponectin mRNA levels and carcass traits showed negative correlations between adiponectin mRNA levels in visceral fat tissue and s.c. fat (r = –0.44, P < 0.05), visceral fat (r = –0.43, P < 0.05), and total body fat (r = –0.42, P < 0.05) in the overall gilt population (Table 3). Within-breed analyses showed a negative correlation between adiponectin mRNA levels measured in s.c. fat and s.c. (r = –0.78, P < 0.01), visceral (r = –0.73, P < 0.05), and total body (r = –0.73, P < 0.05) fat weights for the HL gilts only (Table 3). In visceral fat tissue, there was also a significant negative correlation between adiponectin mRNA abundance and s.c. (r = –0.63, P < 0.05) and total body (r = –0.67, P < 0.05) fat weights, whereas there was a nonsignificant correlation with visceral fat (r = –0.57, P = 0.067) in the UM gilts. In the same tissue, the HL gilts showed a nonsignificant correlation between adiponectin mRNA levels and visceral fat (r = –0.54, P = 0.074). Finally, no correlation could be found between adiponectin mRNA levels in both tissues and plasma concentrations of insulin or glucose (data not shown).
Expression of Pig Adiponectin Receptors 1 and 2 mRNA in Subcutaneous and Visceral Fat
Pig adipoR1 and adipoR2 mRNA levels were also quantified by real-time PCR in s.c. and visceral fat tissues from the three breeds. In the s.c. fat tissue, there were no significant breed differences for either adipoR1 or adipoR2 mRNA levels (Figure 4A). In the visceral fat tissue (Figure 4B), the LW gilts had higher adipoR1 and adipoR2 mRNA levels compared with the UM gilts (P < 0.05). Correlation analyses performed in the overall gilts population showed visceral fat adipoR2 mRNA levels to be negatively correlated with s.c. (r = –0.57, P < 0.01), visceral (r = –0.46, P < 0.05), and total body (r = –0.56, P < 0.01) fat (Table 4). There were no other correlations between carcass traits and adipoR1 and adipoR2 mRNA levels in the overall gilt population. Within-breed analyses of adipoR1 mRNA levels in visceral fat showed significant negative correlation with visceral fat (r = –0.72, P < 0.05), whereas a tendency was observed with s.c. fat weight (r = –0.58, P = 0.10), and a nonsignficant correlation was noted with total body (r = –0.60, P = 0.060) fat weight for the LW gilts only (Table 4). Within-breed analyses of adipoR2 mRNA levels in visceral fat showed a significant negative correlation with s.c. (r = –0.68, P < 0.05) and total body (r = –0.71, P < 0.05) fat weights, whereas there was a nonsignificant correlation with visceral fat (r = –0.63, P = 0.070) for the LW gilts (Table 4). For the UM gilts, negative correlations also were observed between adipoR2 mRNA levels in visceral fat and s.c. fat (r = –0.66, P < 0.05) and total body fat weights (r = –0.70, P < 0.05; Table 4).
View larger version (17K):
[in this window]
[in a new window]
|
Figure 4. Relative abundance of adipoR1 and adipoR2 mRNA in s.c. (A) and visceral (B) adipose tissues. Values (±SEM) are expressed as mRNA quantities relative to the UM reference group, which was given a value of one. Ham Line = HL (n = 10), Large White = LW, (n = 10), and Upton-Meishan = UM (n = 10). The asterisks indicate that means differed, P < 0.05.
|
|
View this table:
[in this window]
[in a new window]
|
Table 4. Overall and within-breed correlation coefficients between adipoR1 and adipoR2 mRNA levels and carcass traitsa
|
|
Quantification of Adiponectin and Adiponectin Receptors 1 and 2 in Porcine Stromal-Vascular Cell Culture
Expression of adiponectin, adipoR1, and adipoR2 mRNA levels were determined during differentiation of porcine SV cells in the presence of ITS and were compared with ITS + leptin or ITS + TNF treatments. A significant time-independent decrease in adiponectin mRNA levels was observed when SV cells were incubated with 100 ng/mL of recombinant porcine leptin (Figure 5A; P < 0.05). Porcine leptin did not significantly influence adipoR1 and adipoR2 mRNA levels (Figure 5B and C). A significant time-dependent decrease of adiponectin mRNA levels could be observed when SV cells were treated with 10 ng/mL of recombinant human TNF (Figure 5A; P < 0.001). A significant time-independent decrease was also observed for adipoR2 mRNA levels in SV cells treated with human TNF (Figure 5C; P < 0.001). In contrast, mRNA levels of adipoR1 were not influenced by TNF treatment (Figure 5B). We next examined the effects of TNF and recombinant porcine leptin on the expression of aP2, which is a well-known marker of adipocyte differentiation. Lower mRNA levels of aP2 were observed in SV cells treated with ITS + leptin over a 96-h period, and this effect was more pronounced with ITS + TNF (Figure 5D). Compared with the ITS-treated cells, decreases in lipid accumulation were observed in SV cells treated with ITS + TNF, as revealed by Oil Red O staining after 24, 48, and 96 h exposure to TNF (data not shown).
|
Discussion
|
|---|
To our knowledge, this is the first report of the cloning and characterization of pig adipoR1 and adipoR2. The deduced porcine adipoR1 protein sequence shares a high level of similarity with human and mouse orthologues (Yamauchi et al., 2003), and the deduced porcine adipoR2 protein sequence is identical with the human adipoR2 and nearly identical with the mouse sequence (Yamauchi et al., 2003). The main feature distinguishing the pig adipoR1 protein from human and mouse proteins is a hypervariable region between AA 39 and 51 relative to the human protein sequence. In this hypervariable region, three threonine residues are found in the porcine sequence that are not present in the human and mouse counterparts. Prediction of putative Thr phosphorylation sites, using the PhosphoBase database (Kreegipuu et al., 1999) and the ELM motif search (http://ca.expasy.org/tools/), revealed that Thr16 (relative to the porcine sequence) may be phosphorylated by adenosine cyclic 3',5'-phosphate protein kinase. Because AA phosphorylation is central to signal transduction pathways, this species discrepancy may have functional relevance to adiponectin signal transduction.
Adiponectin mRNA was abundantly expressed in s.c. and visceral fat tissues, consistent with human (Maeda et al., 1996) and mouse (Hu et al., 1996) tissue distributions. In the current study, we also detected, for the first time, adiponectin mRNA in pig skeletal muscle tissue, derived from i.m. fat cells or from muscle cells, as it was previously reported that adiponectin mRNA expression is inducible in human myotubes (Staiger et al., 2003). Adiponectin receptors mRNA is ubiquitously expressed in pig tissues, as are the receptors’ orthologues in human and mouse species (Yamauchi et al., 2003). The pig R1 subtype is abundantly expressed in skeletal muscle, visceral fat, and s.c. fat tissues, whereas the R2 isoform is predominantly expressed in liver, heart, skeletal muscle, visceral fat, and s.c. fat tissues. In was recently reported that adiponectin receptors also are expressed in 3T3-L1 adipocytes (Fasshauer et al., 2004), but this is the first time that adipoR1 and adipoR2 mRNA expression has been demonstrated in visceral and s.c. fat tissues. The presence of adipoR1 and adipoR2 in porcine fat tissues suggests that adiponectin may have a paracrine role in fat cells. Alternatively, circulating adiponectin may act directly on these cells to suppress lipogenesis. Such a role was recently reported by Jacobi et al. (2004), who found that adiponectin directly affects isolated pig adipocytes to attenuate lipogenesis. In the current study, we show for the first time that the mRNA of both adiponectin receptors is expressed in the ovary and uterus, thereby suggesting that adiponectin may have effects on pig reproductive function as well.
In the present study, we also found that the porcine adiponectin mRNA levels are higher in visceral than in s.c. fat. These results are in accordance with those of Altomonte et al. (2003), where lean rats showed higher levels of adiponectin mRNA in visceral fat than in s.c. adipose tissue. These authors suggested that visceral fat might be a predominant contributor to plasma adiponectin. Few human studies have examined adiponectin mRNA levels in various fat depots, and yet these results are inconsistent, showing either no difference (Yang et al., 2003) or lower levels of adiponectin mRNA in omental (Fisher et al., 2002) or visceral fat (Lihn et al., 2004) compared to s.c. adipose. Interestingly, we observed that the highest visceral/s.c. fat tissue ratios of adiponectin mRNA levels were found in UM, followed by LW and HL. These results show that visceral adipocytes from fatter pigs express higher levels of adiponectin mRNA in visceral fat compared with adipocytes from s.c. fat tissues. In the current study, we only had access to females of the three genotypes used. Because other studies have reported gender differences in circulating adiponectin (Arita et al., 1999; Combs et al., 2003), care should be taken in extrapolating these results to both genders. Despite the fact that we observed higher levels of adiponectin mRNA in porcine visceral fat, the s.c. fat depot may also contribute substantially more adiponectin to the circulation than visceral fat because s.c. fat depots (which includes intermuscular fat depots) comprise approximately 92% of the total fat mass in our pig sample.
The Meishan pigs are recognized for their slower growth and higher backfat thickness than Occidental breeds (Bonneau et al., 1990). In the present study, UM gilts, although a crossbreed between Meishan and an Occidental breed, had greater visceral, s.c., and total body fat than HL and LW gilts. These UM gilts also had lower adiponectin mRNA levels in their s.c. fat tissue compared with the leaner HL gilts. Similarly, Jacobi et al. (2004) showed that there was no difference in pig adiponectin protein concentrations between fat and lean genotype at 28 and 56 d of age, whereas significant differences were reported at 90 and 165 d of age. Adiponectin mRNA levels and its plasma concentration are also decreased with developing obesity in rats and human subjects, but the mechanism underlying the decrease in adiponectin production is not fully understood (Hu et al., 1996; Arita et al., 1999; Milan et al., 2002). Our data clearly show that adiponectin mRNA levels in s.c. and visceral fat tissues are not associated with circulating insulin and glucose concentrations in pigs. Although most studies in humans and rodents report significant associations between adiponectin levels and insulin resistance (Weyer et al., 2001; Altomonte et al., 2003), a more limited role for adiponectin on insulin action has been suggested by other reports. For example, Silha et al. (2003) demonstrated that insulin resistance was more closely related to circulating resistin and leptin than to adiponectin levels. Other studies have also reported that adiponectin deficiency can results in no insulin resistance or glucose intolerance (Ma et al., 2002; Silha et al., 2004).
In this study, the s.c., visceral, and total body fat depots weights were negatively correlated with adiponectin mRNA levels from visceral adipose tissue, but not from s.c. fat tissue. Milan et al. (2002) previously reported that adiponectin expression was significantly lower in visceral fat tissue from genetically obese vs. lean rats. Consistent with our results, these authors could not find such differences when s.c. fat tissues were compared from the same animals. Interestingly, previously reported studies (Cnop et al., 2003; Park et al., 2004) have suggested that adiponectin expression is determined predominantly by the visceral fat compartment. Within-breed correlation analyses revealed that the association between adiponectin mRNA levels in visceral fat and the various fat depots weights was due mostly to the high negative associations found in the UM breed. No such associations could be made in the leaner breeds of pigs (HL and LW), although this lack of statistical significance might be due to the low number of animals used for each breed. These results suggest that associations between pig adiponectin mRNA levels in visceral fat and pig body fat mass can only be made when pigs have reached higher levels of body fat. Similarly, a study performed on normal-weight and obese women showed that the association found between circulating adiponectin and body fat mass became stronger after an adjustment was made for body fat mass (i.e., adiponectin/body fat mass; Matsubara et al., 2002). Because most associations reported herein were found between adiponectin mRNA levels in visceral fat tissue and the s.c., visceral, and total body fat depots, the significance of the high negative correlations between adiponectin mRNA levels in s.c. fat tissue and the various fat depots weights in the HL gilts only remains unclear to us.
In the current study, we provide the first evidence that pig adipoR1 and adipoR2 mRNA is less expressed in fat pigs when compared with lean breeds, and that this effect can only be detected in visceral fat tissue. Similar results were reported by Tsuchida et al. (2004), who found lower mRNA levels of adipoR1 and adipoR2 in white adipose tissue of ob/ob mice, a model of insulin resistance and obesity. Moreover, these authors reported that adipoR1 and adipoR2 mRNA levels in ob/ob mice were 31.9 and 7.9% of those from the wild-type mice, respectively. The same observation can be made in pigs, where adipoR1 and adipoR2 mRNA levels in UM fat gilts are 66 and 37% of those found in lean LW pigs. Tsuchida et al. (2004) reported that the expression of adipoR1 and adipoR2 is inversely correlated with plasma insulin levels. In contrast, human adiponectin receptors are not altered in muscle of type 2 diabetic patients (Debard et al., 2004), and insulin does not affect adipoR1 and adipoR2 gene expression in 3T3-L1 adipocytes (Fasshauer et al. 2004). In the current study, it seems that pig adipoR1 and adipoR2 mRNA levels are controlled by factors other than insulin and glucose because fasting plasma concentrations of glucose and insulin were similar in all three breeds. Interestingly, Mersmann et al. (1982) previously reported that genetically obese pigs do not manifest their propensity to obesity through modifications in circulating glucose and insulin. Moreover, Wangness et al. (1977) showed that obese pigs are not hyperinsulinemic and have fasting glucose concentrations that are similar to those found in lean pigs.
Correlation analyses performed on the whole population revealed that pig obesity is associated with decreased expression of adipoR2 mRNA but not of adipoR1. Our results also suggest that adipoR2 expression is mainly modulated in the visceral fat tissue, which is also the predominant fat compartment for the control of adiponectin expression (Halleux et al., 2001). A lower expression of adipoR2 mRNA may lead to a decrease in adiponectin binding and thus results in decreased adiponectin effects in visceral adipose tissues. Pig adipoR1 mRNA, which seems to be less affected by the development of obesity, might be able to compensate for decreased expression of adipoR2 in visceral fat, and thus be able to mediate adiponectin effects in fat tissues. There were no correlations between adipoR2 mRNA levels measured in visceral fat tissue and the various fat compartments weights in the HL breed. These discrepancies further suggest that adiponectin receptor gene expression can be modulated differently in adipocytes according to the breed of pigs. It may also suggest that adiponectin effects might be oriented towards various target tissues, such as the liver, muscles, and adipocytes according to the breed of pigs. Thus, it will be of interest to further study breed effects on adiponectin receptors mRNA levels in muscle and liver tissues.
Recent evidence suggests that TNF may be involved in the modulation of lipid metabolism. Indeed, elevated levels of TNF mRNA and protein have been found in adipose tissues of obese rodents, humans, and pigs (Hotamisligil 1999; Chen et al., 2004). Moreover, TNF was the first adipocytokine proposed to represent a molecular link between obesity and insulin resistance (Moller 2000). In the current study, decreases in lipid accumulation and in aP2 mRNA levels were observed in SV cells treated with ITS + TNF. These results agree with those of Rosenbaum and Greenberg (1998), who reported a 200% decrease of aP2 mRNA levels after 48 and 72 h of treatment with TNF on 3T3-L1 adipocytes. Ruan et al. (2002) recently reported that TNF can downregulate many adipocyte-abundant genes that are critical for insulin responsiveness. Because of the known association between adiponectin expression and insulin resistance, we hypothesize that TNF might also modulate adiponectin and adiponectin receptors mRNA levels in SV cells.
In our in vitro system, TNF significantly decreased adiponectin mRNA levels in porcine SV cells in a time-dependent manner. This result agrees with those of Fasshauer et al. (2002), who showed that the expression and secretion of adiponectin from 3T3-L1 adipocytes was significantly decreased by TNF (Fasshauer et al., 2002), and suggested that TNF might have an active role for the decreased adiponectin production in obesity. In the current study, we demonstrate, for the first time, that adipoR2, but not adipoR1, is downregulated by TNF. These results suggest that TNF might mediate its adiponectin-reducing effect via a downregulation of adipoR2 mRNA levels. Previous data suggested that TNF has no significant effect on adipoR2 mRNA levels in 3T3-L1 cell culture (Fasshauer et al., 2004). In this later study, a steady decrease in adipoR2 mRNA levels could be seen through the 4-h incubation period with TNF. We believe that longer incubation periods with TNF, such as those used in the current study, are required to document significant decreases of adipoR2 mRNA levels in fat cells. In the current study, we also demonstrated that adiponectin mRNA levels are significantly decreased when SV cells are incubated with recombinant porcine leptin. Similarly, adiponectin mRNA levels were suppressed by expression of leptin transgene in the hypothalamus of wild type and ob/ob mice (Ueno et al., 2004). In contrast, in was recently reported that leptin directly stimulates adiponectin mRNA and protein levels from 3T3-F442A adipocytes (Delporte et al., 2004). Finally, acute leptin treatment did not modify plasma adiponectin levels in humans (Gravila et al., 2003). Because of the conflicting results reported thus far, further studies will be needed to elucidate the effects of leptin on adiponectin expression.
|
Implications
|
|---|
The present study demonstrates the presence of adiponectin receptors mRNA in pig tissues. We also report that adiponectin and levels of adiponectin receptor 2, but not adiponectin receptor 1, mRNA are modulated in the visceral fat tissues, thereby suggesting an important role of central obesity in pig adiponectin expression. Furthermore, we showed that tumor necrosis factor- may mediate this adiponectin-reducing effect through a downregulation of adiponectin receptor R2, but not adiponectin receptor R1, mRNA levels. Associations between adiponectin, tumor necrosis factor-, and obesity in the pig indicate the utility of the pig as a research model for human studies. Increased understanding of the mechanisms regulating adiponectin expression in the pig will contribute to the development of approaches to treat obesity in humans and to control fat deposition in pigs.
|
Footnotes
|
|---|
1 This work was supported by the National Sciences and Engineering Research Council of Canada Strategic Grant No. 246154 to B. D. Murphy and M. F. Palin, Genetiporc Inc. (St-Bernard, QC, Canada), Agriculture and Agri-Food Canada, and Hypor (formerly Genex Swine Group, Regina, SK, Canada). E. Lord is supported by a FPPQ fellowship. 
2 The authors are grateful to R. Charest for technical assistance, the staff of the Swine Complex for animal care, and S. Méthot for statistical analysis. Lennoxville Dairy and Swine R&D Centre Contribution No. 845.
3 Correspondence: P.O. Box 90, 2000 Rte. 108 E. (phone: 819-565-9174, ext. 207; fax: 819-564-5507; e-mail: palinmf{at}agr.gc.ca).
Received for publication August 19, 2004.
Accepted for publication December 7, 2004.
|
Literature Cited
|
|---|
Altomonte, J., S. Harbaran, A. Richter, and H. Dong. 2003. Fat depot-specific expression of adiponectin is impaired in Zucker fatty rats. Metabolism 52:958–963.[Medline]
Applied Biosystems. 1997. User Bulletin #2: ABI PRISM 7700 Sequence Detection System. Applied Biosystems, Foster City, CA.
Arita, Y., S. Kihara, N. Ouchi, M. Takahashi, K. Maeda, J. Miyagawa, K. Hotta, I. Shimomura, T. Nakamura, K. Miyaoka, H. Kuriyama, M. Nishida, S. Yamashita, K. Okubo, K. Matsubara, M. Muraguchi, Y. Ohmoto, T. Funahashi, and Y. Matsuzawa. 1999. Paradoxical decrease of an adipose-specific protein, adiponectin, in obesity. Biochem. Biophys. Res. Commun. 257:79–83.[Medline]
Bonneau, M., J. Mourot, J. Noblet, L. Lefaucheur, and J. P. Bidanel. 1990. Tissue development in meishan pigs: Muscle and fat development and metabolism and growth hormone regulation by somatotrophic hormone. Pages 201–213 in Proc. Chinese Pig Symp. M. Molenat and C. Legault, ed. Toulouse, France.
Boone, C., F. Gregoire, and C. Remacle. 2000. Culture of porcine stromal-vascular cells in serum-free medium: Differential action of various hormonal agents on adipose conversion. J. Anim. Sci. 78: 885–895.[Abstract/Free Full Text]
CCAC. 1993. Guide to the Care and Use of Experimental Animals. Vol. 1. 2nd ed. Can. Counc. Anim. Care, Ontario, Canada.
Chen, X. D., T. Lei, T. Xia, L. Gan, and Z. Q. Yang. 2004. Increased expression of resistin and tumour necrosis factor-alpha in pig adipose tissue as well as effect of feeding treatment on resistin and cAMP pathway. Diabetes Obes. Metab. 6:271–279.[Medline]
Cnop, M., P. J. Havel, K. M. Utzschneider, D. B. Carr, M. K. Sinha, E. J. Boyko, B. M. Retzlaff, R. H. Knopp, J. D. Brunzell, and S. E. Kahn. 2003. Relationship of adiponectin to body fat distribution, insulin sensitivity and plasma lipoproteins: Evidence for independent roles of age and sex. Diabetelogia 46:459–469.[Medline]
Combs, T. P., A. H. Berg, M. W. Rajala, S. Klebanov, P. Iyengar, J. C. Jimenez-Chillaron, M. E. Patti, S. L. Klein, R. S. Weinstein, and P. E. Scherer. 2003. Sexual differentiation, pregnancy, calorie restriction, and aging affect the adipocyte-specific secretory protein adiponectin. Diabetes 52:268–276.[Abstract/Free Full Text]
Coppack, S. W. 2001. Pro-inflammatory cytokines and adipose tissue. Proc. Nutr. Soc. 60:349–356.[Medline]
Debard, C., M. Laville, V. Berbe, E. Loizon, C. Guillet, B. Morio-Liondore, Y. Boirie, and H. Vidal. 2004. Expression of key genes of fatty acid oxidation, including adiponectin receptors, in skeletal muscle of type 2 diabetic patients. Diabetologia 47:917–925.[Medline]
Delporte, M.-L., S. Ait El Mkadem, M. Quisquater, and S. M. Brichard. 2004. Leptin treatment markedly increased plasma adiponectin, but barely decreased plasma resistin of ob/ob mice. Am. J. Physiol. Endocrinol. Metab. 287:E446–E453.[Abstract/Free Full Text]
Ding, S. T., R. L. McNeel, and H. J. Mersmann. 1999. Expression of porcine adipocyte transcripts: Tissue distribution and differentiation in vitro and in vivo. Comp. Biochem. Physiol. B. Biochem. Mol. Biol. 123:307–318.[Medline]
Fasshauer, M., J. Klein, S. Kralisch, M. Klier, U. Lossner, M. Bluher, and R. Paschke. 2004. Growth hormone is a positive regulator of adiponectin receptor 2 in 3T3-L1 adipocytes. FEBS Lett. 558:27–32.[Medline]
Fasshauer, M., J. Klein, S. Neumann, M. Eszlinger, and R. Paschke. 2002. Hormonal regulation of adiponectin gene expression in 3T3-L1 adipocytes. Biochem. Biophys. Res. Commun. 290:1084–1089.[Medline]
Fisher, F. M., P. G. McTernan, G. Valsamakis, R. Chetty, A. L. Harte, A. J. Anwar, J. Starcynski, J. Crocker, A. H. Barnett, C. L. McTernan, and S. Kumar. 2002. Differences in adiponectin protein expression: Effect of fat depots and type 2 diabetic status. Horm. Metab. Res. 34:650–654.[Medline]
Gavrila, A., J. L. Chan, N. Yiannakouris, M. Kontogianni, L. C. Miller, C. Orlova, and C. S. Mantzoros. 2003. Serum adiponectin levels are inversely associated with overall and central fat distribution but are not directly regulated by acute fasting or leptin administration in humans: Cross-sectional and interventional studies. J. Clin. Endocrinol. Metab. 88:4823–4831.[Abstract/Free Full Text]
Halleux, C. M., M. Takahashi, M. L. Delporte, R. Detry, T. Funahashi, Y. Matsuzawa, and S. M. Brichard. 2001. Secretion of adiponectin and regulation of apM1 gene expression in human visceral adipose tissue. Biochem. Biophys. Res. Commun. 288:1102–1107.[Medline]
Hausman, G. J. 1989. The influence of insulin, triiodothyronine (T3) and insulin-like growth factor-I (IGF-1) on the differentiation of preadipocytes in serum-free cultures of pig stromal-vascular cells. J. Anim. Sci. 67:3136–3143.
Hotamisligil, G. S. 1999. The role of TNF and TNF receptors in obesity and insulin resistance. J. Intern. Med. 245:621–625.[Medline]
Hotta, K., T. Funahashi, Y. Arita, M. Takahashi, M. Matsuda, Y. Okamoto, H. Iwahashi, H. Kuriyama, N. Ouchi, K. Maeda, M. Nishida, S. Kihara, N. Sakai, T. Nakajima, K. Hasegawa, M. Muraguchi, Y. Ohmoto, T. Nakamura, S. Yamashita, T. Hanafusa, and Y. Matsuzawa. 2000. Plasma concentrations of a novel, adipose-specific protein, adiponectin, in type 2 diabetic patients. Arterioscler. Thromb. Vasc. Biol. 20:1595–1599.[Abstract/Free Full Text]
Hu, E., P. Liang, and B. M. Spiegelman. 1996. AdipoQ is a novel adipose-specific gene dysregulated in obesity. J. Biol. Chem. 271:10697–10703.[Abstract/Free Full Text]
Jacobi, S. K., K. M. Ajuwon, T. E. Weber, J. L. Kuske, C. J. Dyer, and M. E. Spurlock. 2004. Cloning and expression of porcine adiponectin, and its relationship to adiposity, lipogenesis and the acute phase response. J. Endocrinol. 182:133–144.[Abstract]
Kreegipuu, A., N. Blom, and S. Brunak. 1999. PhosphoBase, a database of phosphorylation sites: Release 2.0. Nucleic Acids Res. 27:237–239.[Abstract/Free Full Text]
Kubota, N., Y. Terauchi, T. Yamauchi, T. Kubota, M. Moroi, J. Matsui, K. Eto, T. Yamashita, J. Kamon, H. Satoh, W. Yano, P. Froguel, R. Nagai, S. Kimura, T. Kadowaki, and T. Noda. 2002. Disruption of adiponectin causes insulin resistance and neointimal formation. J. Biol. Chem. 277:25863–25866.[Abstract/Free Full Text]
Lihn, A. S., J. M. Bruun, G. He, S. B. Pedersen, P. F. Jensen, and B. Richelsen. 2004. Lower expression of adiponectin mRNA in visceral adipose tissue in lean and obese subjects. Mol. Cell. Endocrinol. 219:9–15.[Medline]
Ma, K., A. Cabrero, P. K. Saha, H. Kojima, L. Li, B. H. Chang, A. Paul, and L. Chan. 2002. Increased beta-oxidation but no insulin resistance or glucose intolerance in mice lacking adiponectin. J. Biol. Chem. 277:34658–34661.[Abstract/Free Full Text]
Maeda, K., K. Okubo, I. Shimomura, T. Funahashi, Y. Matsuzawa, and K. Matsubara. 1996. cDNA cloning and expression of a novel adipose specific collagen-like factor, apMI (AdiPose Most abundant Gene transcript 1). Biochem. Biophys. Res. Commun. 221:286–289.[Medline]
Maeda, N., I. Shimomura, K. Kishida, H. Nishizawa, M. Matsuda, H. Nagaretani, N. Furuyama, H. Kondo, M. Takahashi, Y. Arita, R. Komuro, N. Ouchi, S. Kihara, Y. Tochino, K. Okutomi, M. Horie, S. Takeda, T. Aoyama, T. Funahashi, and Y. Matsuzawa. 2002. Diet-induced insulin resistance in mice lacking adiponectin/ACRP30. Nat. Med. 8:731–737.[Medline]
Magun, R., B. M. Burgering, P. J. Coffer, D. Pardasani, Y. Lin, J. Chabot, and A. Sorisky. 1996. Expression of a constitutively activated form of protein kinase B (c-Akt) in 3T3-L1 preadipose cells causes spontaneous differentiation. Endocrinology. 137:3590–3593.[Abstract]
Marcoux, M., J. F. Bernier, and C. Pomar. 2003. Estimation of Canadian and European lean yields and composition of pig carcasses by dual-energy x-ray absorptiometry. Meat Sci. 63:359–365.
Matsubara, M., S. Maruoka, and S. Katayose. 2002. Inverse relationship between plasma adiponectin and leptin concentrations in normal-weight and obese women. Eur. J. Endocrinol. 147:173–180.[Abstract]
Mersmann, H. J., W. G. Pond, and J. T. Yen. 1982. Plasma glucose, insulin and lipids during growth of genetically lean and obese swine. Growth 46:189–198.[Medline]
Milan, G., M. Granzotto, A. Scarda, A. Calcagno, C. Pagano, G. Federspil, and R. Vettor. 2002. Resistin and adiponectin expression in visceral fat of obese rats: Effect of weight loss. Obes. Res. 10:1095–1103.[Medline]
Moller, D. E. 2000. Potential role of TNF-alpha in the pathogenesis of insulin resistance and type 2 diabetes. Trends Endocrinol. Metab. 11:212–217.[Medline]
Park, K. G., K. S. Park, M. J. Kim, H. S. Kim, Y. S. Suh, J. D. Ahn, K. K. Park, Y. C. Chang, and I. K. Lee. 2004. Relationship between serum adiponectin and leptin concentrations and body fat distribution. Diabetes Res. Clin. Pract. 63:135–142.[Medline]
Ramsay, T. G. 2001. Porcine leptin alters insulin inhibition of lipolysis in porcine adipocytes in vitro. J. Anim. Sci. 79:653–657.[Abstract/Free Full Text]
Rosenbaum S. E., and A. S. Greenberg. 1998. The short- and long-term effects of tumor necrosis factor-alpha and BRL 49653 on peroxisome proliferator-activated receptor (PPAR) gamma2 gene expression and other adipocyte genes. Mol. Endocrinol. 12:1150–1160.[Abstract/Free Full Text]
Ruan, H., N. Hacohen, T. R. Golub, L. Van Parijs, and H. F. Lodish. 2002. Tumor necrosis factor-alpha suppresses adipocyte-specific genes and activates expression of preadipocyte genes in 3T3-L1 adipocytes: Nuclear factor-kappaB activation by TNF-alpha is obligatory. Diabetes 51:1319–1336.[Abstract/Free Full Text]
Ruiz-Cortes, Z. T., Y. Martel-Kennes, N. Y. Gevry, B. R. Downey, M. F. Palin, and B. D. Murphy 2003. Biphasic effects of leptin in porcine granulosa cells. Biol. Reprod. 68:789–96.[Abstract/Free Full Text]
Silha, J. V., M. Krsek, J. U. Skrha, P. Sucharda, B. L. Nyomba, and L. J. Murphy. 2003. Plasma resistin, adiponectin and leptin levels in lean and obese subjects: correlations with insulin resistance. Eur. J. Endocrinol. 149:331–335.[Abstract]
Silha, J. V., M. Krsek, J. Skrha, P. Sucharda, B. L. Nyomba, and L. J. Murphy. 2004. Plasma resistin, leptin and adiponectin levels in non-diabetic and diabetic obese subjects. Diabet. Med. 21:497–499.
Staiger, H., C. Kausch, A. Guirguis, M. Weisser, E. Maerker, M. Stumvoll, R. Lammers, F. Machicao, and H. U. Haring. 2003. Induction of adiponectin gene expression in human myotubes by an adiponectin-containing HEK293 cell culture supernatant. Diabetologia 46:956–960.[Medline]
Steppan, C. M., S. T. Bailey, S. Bhat, E. J. Brown, R. R. Banerjee, C. M. Wright, H. R. Patel, R. S. Ahima, and M. A. Lazar. 2001. The hormone resistin links obesity to diabetes. Nature (Lond.) 409:307–312.[Medline]
Tsuchida, A., T. Yamauchi, Y. Ito, Y. Hada, T. Maki, S. Takekawa, J. Kamon, M. Kobayashi, R. Suzuki, K. Hara, N. Kubota, Y. Terauchi, P. Froguel, J. Nakae, M. Kasuga, D. Accili, K. Tobe, K. Ueki, R. Nagai, and T. Kadowaki. 2004. Insulin/Foxo1 pathway regulates expression levels of adiponectin receptors and adiponectin sensitivity. J. Biol. Chem. 279:30817–30822.[Abstract/Free Full Text]
Ueno, N., M. G. Dube, A. Inui, P. S. Kalra, and S. P. Kalra. 2004. Leptin modulates orexigenic effects of ghrelin, attenuates adiponectin and insulin levels, and selectively the dark-phase feeding as revealed by central leptin gene therapy. Endocrinology 145:4176–4184.[Abstract/Free Full Text]
Applied Biosystems. 1997. User Bulletin #2: ABI PRISM 7700 Sequence Detection System. Applied Biosystems, Foster City, CA.
Wangsness, P. J., R. J. Martin, and J. H. Gahagan. 1977. Insulin and growth hormone in lean and obese pigs. Am. J. Physiol. 233:E104–E108.[Medline]
Weyer, C., T. Funahashi, S. Tanaka, K. Hotta, Y. Matsuzawa, R. E. Pratley, and P. A. Tataranni. 2001. Hypoadiponectinemia in obesity and type 2 diabetes: Close association with insulin resistance and hyperinsulinemia. J. Clin. Endocrinol. Metab. 86:1930–1935.[Abstract/Free Full Text]
Yamauchi, T., J. Kamon, H. Waki, Y. Terauchi, N. Kubota, K. Hara, Y. Mori, T. Ide, K. Murakami, N. Tsuboyama-Kasaoka, O. Ezaki, Y. Akanuma, O. Gavrilova, C. Vinson, M. L. Reitman, H. Kagechika, K. Shudo, M. Yoda, Y. Nakano, K. Tobe, R. Nagai, S. Kimura, M. Tomita, P. Froguel, and T. Kadowaki. 2001. The fat-derived hormone adiponectin reverses insulin resistance associated with both lipoatrophy and obesity. Nat. Med. 7:941–946.[Medline]
Yamauchi, T., J. Kamon, Y. Ito, A. Tsuchida, T. Yokomizo, S. Kita, T. Sugiyama, M. Miyagishi, K. Hara, M. Tsunoda, K. Murakami, T. Ohteki, S. Uchida, S. Takekawa, H. Waki, N. H. Tsuno, Y. Shibata, Y. Terauchi, P. Froguel, K. Tobe, S. Koyasu, K. Taira, T. Kitamura, T. Shimizu, R. Nagai, and T. Kadowaki. 2003. Cloning of adiponectin receptors that mediate antidiabetic metabolic effects. Nature (Lond.) 423:762–769.[Medline]
Yang, W. S., M. H. Chen, W. J. Lee, K. C. Lee, C. L. Chao, K. C. Huang, C. L. Chen, T. Y. Tai, and L. M. Chuang. 2003. Adiponectin mRNA levels in the abdominal adipose depots of nondiabetic women. Int. J. Obes. Relat. Metab. Disord. 27:896–900.[Medline]
This article has been cited by other articles:
Top
Abstract
Introduction
