Proinflammatory Cytokine and Chemokine Gene Expression Profiles in Subcutaneous and Visceral Adipose Tissue Depots of Insulin‐Resistant and Insulin‐Sensitive Light Breed Horses

Proinflammatory Cytokine and Chemokine Gene Expression Profiles in Subcutaneous and Visceral... Abbreviations : AIRg acute insulin response to glucose EMS equine metabolic syndrome FSIGTT frequently sampled intravenous glucose tolerance test IR insulin resistant IS insulin sensitive RT‐qPCR real‐time quantitative polymerase chain reaction SC subcutaneous SI insulin sensitivity Local expression of inflammatory molecules by adipose tissue depots is reported to play a central role in the onset of local and systemic insulin resistance in the obesity‐related disease metabolic syndrome in both humans and horses. Equine metabolic syndrome (EMS), an increasingly well‐characterized constellation of clinical findings including obesity (particularly regional adiposity), systemic insulin resistance, hyperinsulinemia, dyslipidemia, and increased risk of endocrinopathic laminitis, has been suggested to represent a state of systemic inflammation similar to that reported in humans. Historically, white adipose tissue was considered a relatively metabolically inert tissue that functioned primarily as an energy storage depot. However, since the discovery and description of the adipose tissue‐derived hormone leptin in 1995, white adipose tissue has been increasingly recognized to be not only highly metabolically active, but also endocrinologically important in regulating metabolism. With the development of obesity, adipose tissue is reported to adopt a distinctly inflammatory phenotype, characterized by increased gene expression and secretion of proinflammatory cytokines (tumor necrosis factor‐α [TNF‐α], interleukin‐1β [IL‐1β], and IL‐6) and chemokines. Proinflammatory cytokine and chemokine expression patterns have been shown to be heterogeneous between adipose tissue depots in humans and rodents, with visceral adipose tissue (omental, retroperitoneal, and mesenteric depots) displaying greater gene expression of these substances both in lean individuals and in the setting of genetic and nutritional obesity. Knowledge of these depot‐specific differences has led to the categorization of omental and truncal obesity as a strong risk factor for atherothrombotic cardiovascular disease in obese humans. Although metabolic risk has been documented for obese ponies with prominent regional adiposity (ie, large nuchal ligament adipose tissue accumulation), the relationship of this risk to metabolic activity of the adipose tissue itself has not been reported in horses. Systemic insulin resistance is a defining characteristic of EMS and appears to be strongly linked to laminitis risk in affected animals. Hyperinsulinemia, a common sequel to peripheral insulin resistance in the EMS patient, recently has been shown to precipitate clinical laminitis in ponies without EMS. Therefore, insulin resistance can be considered both a defining marker of EMS and, possibly, as a result of increased insulin concentration, a central mechanism in the pathophysiology of its most important complication, laminitis. Because adipose tissue‐derived inflammatory molecules are reported to play a central role in the onset of insulin resistance, the study reported here was performed to investigate proinflammatory cytokine and chemokine gene expression in different adipose tissue depots and to determine the relationship of inflammatory gene expression patterns in the different depots to systemic insulin resistance in adult light breed horses. Materials and Methods Experimental Animals Light breed mares owned by The Ohio State University College of Veterinary Medicine and housed at the college teaching and research farm were utilized for this project. All mares were group‐housed in a dry lot and fed timothy hay ad libitum (grown and baled on‐site); forage nutrient analysis was not performed, but all mares were fed identically. No clinical evidence of laminitis was observed in any of the animals used for this study. Experimental procedures were performed according to protocols approved by the OSU Institutional Animal Care and Use Committee in accordance with the NIH Guide for the Care and Use of Laboratory Animals. Insulin‐Modified Frequently Sampled Intravenous Glucose Tolerance Testing (FSIGTT) FSIGTT was performed on 20 mares, 12 of which were initially selected for adipose tissue biopsy specimen collection based on FSIGTT results (a power analysis performed before initiation of the study suggested that the number of animals used in this study was appropriate for the measured variables). All tests were performed in April and June 2008. Testing was performed as reported previously by Pratt et al. Briefly, IV catheters were placed in the right and left external jugular veins of each horse after aseptic preparation and subcutaneous (SC) infiltration of the sites with local anesthetic. One catheter was designated for medication administration, and one was designated for blood collection. The horses were confined in stalls overnight after catheter placement to minimize the effect of stress on test results. Blood initially was collected for baseline measurement of plasma insulin and glucose concentrations, followed by administration of a dose of 50% dextrose (300 mg/kg IV, approximately 300 mL, administered over 2–3 minutes). Completion of this infusion marked time ( t ) = 0. Blood samples (approximately 6–8 mL/time point) were collected in EDTA for measurement of insulin and glucose concentrations at 0, 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 12, 14, 16, and 19 minutes postdextrose infusion. A dose of regular insulin (0.1 IU/kg IV) was administered at t = 20 minutes, and additional blood samples were collected at 22, 24, 26, 30, 35, 40, 50, 60, 70, 80, 90, 100, 120, 150, and 180 minutes postdextrose infusion. After completion of sampling, the catheters were removed and the animals returned to the herd. Insulin concentrations were determined by a commercially available radioimmunoassay validated previously for horses, and glucose concentrations were analyzed with a colorimetric assay. Insulin and glucose data were analyzed by minimal model parameters, and the calculated insulin sensitivity (SI; L/min/mU) was used to categorize each horse as insulin resistant (IR; SI < 1.0 L/min/mU) or insulin sensitive (IS; SI > 1.5 L/min/mU). Based on the SI results, 12 of the 20 tested mares (6 IR and 6 IS) were selected to undergo adipose tissue biopsy specimen collection. Adipose Tissue Biopsy Specimen Collection All biopsy specimen collections were performed in July 2008. Horses were transported to the OSU Veterinary Teaching Hospital, weighed upon arrival, and housed individually in box stalls with access to timothy hay, water, and a trace mineral block ad libitum. A physical examination was performed on each horse upon arrival, and morphometric parameters were recorded (neck circumference, girth circumference, ultrasonographic retroperitoneal fat thickness at the umbilicus). All animals were considered healthy based on results of physical examination, CBC, and serum biochemical analysis performed on the day of arrival. Evidence of chronic pyometra, however, was detected in 1 IR mare during hospitalization, and data from this mare therefore were excluded from the final analysis. An acclimation period of 48 hours was observed before biopsy specimen collection. On the day of biopsy specimen collection, a 14 G IV catheter was aseptically inserted in the left external jugular vein for the purposes of sample collection and induction and maintenance of general anesthesia. A pulmonary artery catheter was aseptically inserted via the right external jugular vein for use in collecting blood gas data for a separate study. A tetanus toxoid booster was administered in the left cervical muscle before biopsy specimen collection, and sulfamethoxazole‐trimethoprim (15 mg/kg PO q12h) and phenylbutazone (1 g PO q12h) were administered to each animal (continued for 5 days after biopsy specimen collection). After sedation with xylazine (0.5 mg/kg IV), general anesthesia was induced with ketamine (2.2 mg/kg IV) and diazepam (0.1 mg/kg IV) and maintained with guiafenesin ‐based total IV anesthesia (guiafenesin 5%; xylazine 0.05%; ketamine 0.2%; 1–2 mL/kg/h). All mares were orotracheally intubated, and positive‐pressure ventilation was initiated and maintained with 100% oxygen for the duration of the procedure (approximately 30 minutes). Incisional biopsy specimens of nuchal crest and tail head adipose tissue were collected (∼3 g tissue) with the animal in lateral recumbency after aseptic preparation of the biopsy specimens sites with chlorhexidine surgical scrub and isopropyl alcohol ; incisions were closed in a single layer with 0 polydioxanone monofilament suture material in a simple continuous pattern. Each horse then was placed on a padded surgical table in dorsal recumbency and moved into a surgical suite; the abdomen was aseptically prepared and draped for ventral midline celiotomy. A small ventral midline abdominal incision was created (∼8 cm in length), and adipose tissue biopsy specimens (∼3 g each) were collected from the margins of the incision (retroperitoneal adipose tissue), the mesocolon of the descending colon (mesenteric adipose tissue), and the omentum. The abdominal incision was closed in 2 layers: the linea alba was apposed with 2 polyglactin 910 in a simple continuous pattern, and the skin was apposed with 0 polydioxanone in a simple continuous pattern. Horses then were allowed to recover from general anesthesia and returned to box stalls. All mares remained hospitalized for 10 days after biopsy specimen collection, during which time they were continuously monitored for incisional and other complications. After this period, all mares were returned to the teaching herd. Sutures were removed manually from the nuchal ligament and tail head incisions 14 days after biopsy specimen collection; sutures were not removed from the abdominal incisions. A portion of each adipose tissue biopsy specimen was flash‐frozen in liquid nitrogen at the time of collection and stored at −80°C for later analysis by real‐time quantitative polymerase chain reaction (RT‐qPCR); the remainder was fixed in neutral buffered formalin and paraffin embedded. RNA Isolation, cDNA Synthesis, and RT‐qPCR Homogenates were made from each adipose tissue sample (1.0–1.5 g) with a tissue disruptor, and total RNA was extracted by a modified Trizol method. cDNA was synthesized from total RNA from each sample (after DNAse treatment ) via reverse transcription (Retroscript ) and stored at −20°C until used for RT‐qPCR analysis. A thermocycler (Roche 480 ) was used to perform RT‐qPCR. Amplification was quantified against external standards using fluorescent format for SYBR Green I as described previously. Primers for TNF‐α, IL‐1β, IL‐6, plasminogen activator inhibitor‐1 (PAI‐1), monocyte chemoattractant protein‐1 (MCP‐1), and the housekeeping genes (β‐actin and β 2 ‐microglobulin) were designed against equine‐specific gene sequences (exon sequences) by computer programs as described previously and were used for amplification of the respective genes of interest. Standard curves for quantification of transcript in adipose tissue samples were generated with serial dilutions of a linearized vector containing an insert of each amplified cDNA fragment. Data Analysis RT‐qPCR data from housekeeping genes (β‐actin and β 2 ‐microglobulin) were entered into a computer program to test each gene's quality as a housekeeping gene for equine adipose tissue. Because both genes were determined to be satisfactory by the program, a geometric mean was obtained from the 2 genes' data in order to generate a normalization factor for gene expression from each adipose tissue sample. Gene expression data from the genes of interest then were normalized using this factor for each individual tissue. Data analysis was performed by a statistical software program. Morphometric and gene expression data were assessed for normality by the Shapiro‐Wilk and D'Agostino and Pearson's omnibus normality tests. Morphometric data were analyzed with a Student's t ‐test; gene expression data (or log transforms of nonnormal data) were subjected to 2‐way analysis of variance with repetition on 1 factor (depot) followed by Bonferroni's posttest. Correlation analyses (Pearson's rank‐correlation test) were performed to identify relationships between morphometric parameters and nuchal ligament cytokine gene expression. Statistical significance was accepted at P < .05. Results Differences in age, body weight, body condition score, neck circumference, girth circumference, retroperitoneal adipose tissue thickness, or basal insulin concentration between the IR and IS groups were not detected ( Table 1 ). Subjectively, the omental adipose tissue depot of all horses appeared grossly small at laparotomy. Of the parameters evaluated, the only statistically significant difference between the groups was peripheral SI (IR group, SI = 0.58 ± 0.31 × 10 −4 L/min/mU, n = 5; IS group, SI = 2.59 ± 1.21 × 10 −4 L/min/mU, n = 6; P = .008). No statistically significant difference was observed in acute insulin response to glucose (AIRg) between the groups (IR, AIRg = 821.8 ± 585.8 mU/L/min; IS, AIRg = 394.2 ± 162.4 mU/L/min; P = .18). No correlation was observed between basal insulin concentration and SI ( P = .28). 1 Morphometric characteristics of insulin‐resistant (IR) and insulin‐sensitive (IS) light breed mares from which adipose tissue biopsies were collected (mean ± SD). Group Age (years) Body Weight (lb) Body Condition Score (1–9) Neck Circumference (cm) Girth Circumference (cm) Retroperitoneal Fat Thickness (cm) Basal Insulin Concentration (mIU/L) IR (n = 5) 16.8 ± 4.5 1,122 ± 56 7.2 ± 0.7 90.1 ± 4.2 186.3 ± 4.3 3.0 ± 0.9 22.9 ± 9.9 IS (n = 6) 10.8 ± 7.2 1,193 ± 133 6.75 ± 1.6 93.2 ± 3.1 189.1 ± 6.4 2.7 ± 1.7 15.5 ± 11.9 a Thickness of the retroperitoneal adipose tissue measured ultrasonographically at the level of the umbilicus with a 7.5 mHz linear array probe. The average of 5 measurements was recorded for each horse. No statistically significant effect of SI status on concentrations of mRNA coding for TNF‐α ( P = .29), IL‐1β ( P = .80), IL‐6 ( P = .93), MCP‐1 ( P = .23), or PAI‐1 ( P = .72) was noted within each adipose tissue depot ( Fig 1 ). However, a significant difference in gene expression among depots was observed, with higher concentrations of IL‐6 ( P = .004) and IL‐1β ( P = .0005) mRNA measured in nuchal ligament adipose tissue compared with other depots ( Fig 2 ). No differences in gene expression of TNF‐α ( P = .97), MCP‐1 ( P = .14), or PAI‐1 ( P = .07) were observed among depots ( Fig 2 ). 1 Gene expression of IL‐1β, IL‐6, TNF‐α, MCP‐1, and PAI‐1 by adipose tissue depot. Data from IR and IS groups have been stratified for each depot. No statistically significant differences in gene expression were observed between IR and IS groups for each depot. Prominent individual variability in gene expression was noted in IR and IS animals. OM, omental; RET, retroperitoneal; NUC, nuchal ligament; MES, mesocolonic; TAIL, tail head; TNF‐α, tumor necrosis factor‐α; IL, interleukin; PAI‐1, plasminogen activator inhibitor‐1; MCP‐1, monocyte chemoattractant protein‐1; IR, insulin resistant; IS, insulin sensitive. 2 Gene expression of IL‐1β, IL‐6, TNF‐α, MCP‐1, and PAI‐1 by adipose tissue depot. Data from IR and IS groups have been pooled for each depot. Expression of IL‐1β and IL‐6 is significantly higher in the nuchal ligament adipose tissue depot (*); no other statistically significant differences were observed. OMEN, omental; RETRO, retroperitoneal; NUCHAL, nuchal ligament; MESEN, mesocolonic; TAIL, tail head; TNF‐α, tumor necrosis factor‐α; IL, interleukin; PAI‐1, plasminogen activator inhibitor‐1; MCP‐1, monocyte chemoattractant protein‐1; IR, insulin resistant; IS, insulin sensitive. There was a significant correlation between the expression of IL‐1β and IL‐6 in the nuchal ligament ( r = 0.76; P = .006). However, there was no statistically significant correlation between IL‐1β or IL‐6 expression and mean neck circumference (IL‐1β: r = 0.24, P = .48; IL‐6: r = 0.31, P = .36), body condition score (IL‐1β: r =− 0.10, P = .76; IL‐6: r = 0.11, P = .74), and body weight (IL‐1β: r = 0.08, P = .82; IL‐6: r = 0.15, P = .64). Discussion The study described here compares proinflammatory gene expression profiles of multiple visceral and SC adipose tissue depots of light breed horses. The results suggest that the visceral adipose tissue depots of the adult horse do not have greater expression of genes encoding inflammatory cytokines when compared with SC adipose tissue depots, a finding that differs substantially from observations made in other species. Omental adipose tissue, which has been reported to be the primary depot undergoing inflammatory signaling in obese humans and laboratory animals, and where fat accumulation reportedly correlates closely with cardiovascular disease risk in obese humans, does not appear to display a greater inflammatory gene expression profile than other adipose tissue depots in the adult horse. EMS has emerged in recent years as a substantial risk factor for development of endocrinopathic laminitis in horses and ponies. Whereas systemic insulin resistance, increased adiposity (particularly regional adiposity), hyperinsulinemia, and dyslipidemia characterize the syndrome as it is currently accepted, only insulin resistance has thus far been identified as an independent risk factor for laminitis and directly involved in its pathogenesis. The horses used in this study displayed no physical evidence of prior bouts of laminitis; neither did they have prominent nuchal ligament adipose tissue deposits. These findings suggest that these subjects should not be categorized as having EMS per se. However, because insulin resistance may be the component of EMS most directly associated with laminitis risk, its relationship to proinflammatory cytokine and chemokine gene expression within the adipose tissue was investigated. In humans and experimental rodent models of obesity, adipose tissue (particularly visceral depots) expresses higher concentrations of TNF‐α, IL‐1β, IL‐6, and other proinflammatory cytokines and chemokines in peripherally IR individuals than in those that are IS. Interestingly, the data from the present study do not identify any differences in adipose tissue inflammatory gene expression between depots from IR and those from IS horses of similar body condition scores (all animals being overweight). However, differences in IL‐1β and IL‐6 gene expression among adipose tissue depots were statistically significant, suggesting that the nuchal crest is a more active depot. The lack of an observed effect of peripheral SI on proinflammatory gene expression within a specific depot suggests that the differences noted among depots may be associated with other factors related to obesity. An important confounder in the interpretation of these results is the fact that all of the horses sampled were either overweight or obese according to a widely used body condition scoring system. In several studies of horses, investigators have demonstrated positive correlations between body condition score and peripheral insulin resistance, plasma basal insulin concentration, plasma leptin concentration, proinflammatory cytokine gene expression, and protein concentration in peripheral blood, and risk of laminitis. Negative correlations have been suggested between body condition score and plasma adiponectin concentration in horses, a finding in keeping with this adipokine's reported anti‐inflammatory and cardioprotective value in the setting of human metabolic syndrome. In humans, it is well recognized that obese individuals often are IR. In fact, body mass index and insulin resistance each have been interpreted as proxies for the other in certain assessments of cardiometabolic risk. However, the existence of peripherally IS obese individuals is difficult to explain in this setting. McLaughlin and colleagues recently reported that although, predictably, obesity was more prevalent in a cohort of IR individuals (34%) than lean body composition (16%), obesity and insulin resistance did not make equivalent contributions to combined cardiovascular disease risk in their risk‐assessment model. Their data reinforce the concept that although body mass index and insulin resistance are related, they are not synonymous and contribute independently to cardiovascular risk. Obesity and insulin resistance clearly are correlated in humans and horses, but their respective contributions to subsequent pathology cannot be assumed to be equal. Furthermore, the recent description of a “metabolically healthy but obese” phenotype in several human study populations, in which obesity is notably not accompanied by insulin resistance, hyperglycemia, dyslipidemia, or an increased risk of cardiovascular disease, suggests uncoupling of obesity and metabolic risk in a substantial proportion of individuals. For example, a recent large survey of humans identified 31.7% of obese adults as “metabolically healthy” and 23.5% of lean adults as “metabolically abnormal.” Additionally, very recent work in mice identifies a candidate gene, Brd2, as a molecular link between obesity and insulin resistance. Brd2 knockout mice develop severe obesity and hyperinsulinemia, but they display enhanced glucose tolerance, decreased adipose tissue macrophage infiltration, and increased plasma concentrations of adiponectin. Furthermore, these mice displayed evidence of enhanced expression of proinflammatory cytokines that appeared unrelated to their systemic SI or the degree of adipose tissue macrophage infiltration. Thus, examples from multiple species have weakened the assertion that obesity and insulin resistance are inexorably related. This information may in part explain why a relationship between insulin resistance and gene expression was not identified in this study and why evidence of increased nuchal ligament inflammatory mediator expression was noted in all mares. Therefore, adiposity may factor more importantly in EMS and endocrinopathic laminitis risk than solely through its association with insulin resistance. To further test this possibility, lean cohorts of animals should be included in similar studies in the future to assess whether inflammatory signaling occurs with obesity in the horse regardless of SI. University‐owned mares were an attractive source population for the study reported here because of their ready availability and standardized, uniform management (feeding, housing, and preventive medicine). However, sex influences on both peripheral SI and biological behavior of the adipose tissue have been noted in horses and humans. Additionally, data exist to support the contention that mares have an increased risk of (all‐cause) laminitis than do stallions or geldings, implicating an influence of sex on the most concerning complication of equine obesity. We recognize this influence as a confounder to the interpretation of our results and acknowledge that the data reported here may not be representative of what might be noted in a population of horses including both sexes (intact and altered). An important finding of this study, that nuchal crest fat appears to be the most reactive fat depot (with respect to inflammatory signaling) in the horse, complements clinical findings of previous investigators who have identified the accumulation of nuchal crest adipose tissue as a risk factor for laminitis associated with equine obesity (global, regional, or both). Frank et al reported that obese horses with insulin resistance had greater mean neck circumference scores than nonobese mares, relating this morphologic change with the SI status and body condition of the horse. Accumulation of nuchal ligament adipose tissue has further been shown to have predictive value in assessing risk of pasture‐associated laminitis in multiple cohorts of ponies, with ponies that have larger neck crests being at greater risk for developing the condition. Thus, our finding of unique biological behavior of nuchal crest fat when compared with other depots supports an accumulating body of evidence that the morphology of the nuchal crest changes in the setting of EMS and is correlated with clinical signs of the disease. In conclusion, the current study does not support a direct relationship between the degree of insulin resistance and inflammatory mediator gene expression by adipose tissue in chronically obese horses, suggesting that other factors besides insulin resistance may mark risk for laminar injury in the obese horse. Additionally, the results of the current study indicate that the nuchal ligament adipose depot, rather than visceral adipose depots, may be more important pathophysiologically in the horse. This finding not only fits well with the clinical case at risk of laminitis with enlarged nuchal adipose deposits (ie, a “cresty neck”), but also indicates that it may be possible to focus future adipobiology studies in horses on nuchal ligament tissue biopsy specimens (versus obtaining abdominal or tail head samples). Additional study will be required to determine the role of the adipose tissue in EMS, including the link between changes in the adipose tissue and laminitis and its potential as a therapeutic target in animals at risk of EMS‐associated laminitis. Footnotes a Abbocath 14 G, Abbott Medical Health, Abbott Park, IL b Dextrose 50%, Vedco, St Joseph, MO c BD Vacutainer K 2 EDTA tubes, BD (Becton Dickinson), Franklin Lakes, NJ d Humulin R, Eli Lily and Company, Indianapolis, IN e Coat‐A‐Count insulin RIA, Siemens Medical Solutions Diagnostics, Los Angeles, CA f Thermo‐Trace Ltd, Melbourne, Australia g Tetanus toxoid, Fort Dodge Animal Health/Wyeth, Madison, NJ h Sulfamethoxazole‐trimethoprim 960 mg tablets, Mutual Pharmaceutical Co Inc, Philadelphia, PA i Phenylzone Paste, Schering Plough Animal Health, Union, NJ j Xylazine HCl, 100 mg/mL, IVX Animal Health Inc, St Joseph, MO k Ketaset, Fort Dodge Animal Health/Wyeth l Diazepam Inj, 5 mg/mL, Abbott Laboratories m Guifenesin 5%, Butler Animal Health Supply, Dublin, OH n Chlorhexidine 2% surgical scrub, First Priority Inc, Elgin, IL o Isopropyl alcohol 70%, Butler Animal Health Supply p PDS, Ethicon Inc, Somerville, NJ q Vicryl, Ethicon Inc r Tissue‐Ruptor, Qiagen, Valencia, CA s DNAse I, Thermo Fisher Scientific Inc, Rockford, IL t Retroscript, Ambion Inc, Austin, TX u Roche 480, Roche Applied Science, Indianapolis, IN v TOPO TA Cloning Kit, Invitrogen, Carlsbad, CA w geNorm, Ghent University, Ghent, Belgium x GraphPad Prism 5, GraphPad Software, La Jolla, CA Acknowledgments This work was supported by a grant from the American Quarter Horse Association. The authors acknowledge the following individuals for their assistance with this project: Dr Nicholas Frank and Sarah Elliot (University of Tennessee) for provision of the RNA extraction protocol; Drs Turi Aarnes and Yukie Ueyama for anesthesia support; Drs Britta Leise, Rafael Faleiros, Jarred Williams, Ms Cailing Yin, and Drs Kelly Welin and Trina Westerman for assistance with biopsy specimen collection; and Ms Mauria Watts for critical review of the manuscript. http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Journal of Veterinary Internal Medicine Wiley

Proinflammatory Cytokine and Chemokine Gene Expression Profiles in Subcutaneous and Visceral Adipose Tissue Depots of Insulin‐Resistant and Insulin‐Sensitive Light Breed Horses

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Abstract

Abbreviations : AIRg acute insulin response to glucose EMS equine metabolic syndrome FSIGTT frequently sampled intravenous glucose tolerance test IR insulin resistant IS insulin sensitive RT‐qPCR real‐time quantitative polymerase chain reaction SC subcutaneous SI insulin sensitivity Local expression of inflammatory molecules by adipose tissue depots is reported to play a central role in the onset of local and systemic insulin resistance in the obesity‐related disease metabolic syndrome in both humans and horses. Equine metabolic syndrome (EMS), an increasingly well‐characterized constellation of clinical findings including obesity (particularly regional adiposity), systemic insulin resistance, hyperinsulinemia, dyslipidemia, and increased risk of endocrinopathic laminitis, has been suggested to represent a state of systemic inflammation similar to that reported in humans. Historically, white adipose tissue was considered a relatively metabolically inert tissue that functioned primarily as an energy storage depot. However, since the discovery and description of the adipose tissue‐derived hormone leptin in 1995, white adipose tissue has been increasingly recognized to be not only highly metabolically active, but also endocrinologically important in regulating metabolism. With the development of obesity, adipose tissue is reported to adopt a distinctly inflammatory phenotype, characterized by increased gene expression and secretion of proinflammatory cytokines (tumor necrosis factor‐α [TNF‐α], interleukin‐1β [IL‐1β], and IL‐6) and chemokines. Proinflammatory cytokine and chemokine expression patterns have been shown to be heterogeneous between adipose tissue depots in humans and rodents, with visceral adipose tissue (omental, retroperitoneal, and mesenteric depots) displaying greater gene expression of these substances both in lean individuals and in the setting of genetic and nutritional obesity. Knowledge of these depot‐specific differences has led to the categorization of omental and truncal obesity as a strong risk factor for atherothrombotic cardiovascular disease in obese humans. Although metabolic risk has been documented for obese ponies with prominent regional adiposity (ie, large nuchal ligament adipose tissue accumulation), the relationship of this risk to metabolic activity of the adipose tissue itself has not been reported in horses. Systemic insulin resistance is a defining characteristic of EMS and appears to be strongly linked to laminitis risk in affected animals. Hyperinsulinemia, a common sequel to peripheral insulin resistance in the EMS patient, recently has been shown to precipitate clinical laminitis in ponies without EMS. Therefore, insulin resistance can be considered both a defining marker of EMS and, possibly, as a result of increased insulin concentration, a central mechanism in the pathophysiology of its most important complication, laminitis. Because adipose tissue‐derived inflammatory molecules are reported to play a central role in the onset of insulin resistance, the study reported here was performed to investigate proinflammatory cytokine and chemokine gene expression in different adipose tissue depots and to determine the relationship of inflammatory gene expression patterns in the different depots to systemic insulin resistance in adult light breed horses. Materials and Methods Experimental Animals Light breed mares owned by The Ohio State University College of Veterinary Medicine and housed at the college teaching and research farm were utilized for this project. All mares were group‐housed in a dry lot and fed timothy hay ad libitum (grown and baled on‐site); forage nutrient analysis was not performed, but all mares were fed identically. No clinical evidence of laminitis was observed in any of the animals used for this study. Experimental procedures were performed according to protocols approved by the OSU Institutional Animal Care and Use Committee in accordance with the NIH Guide for the Care and Use of Laboratory Animals. Insulin‐Modified Frequently Sampled Intravenous Glucose Tolerance Testing (FSIGTT) FSIGTT was performed on 20 mares, 12 of which were initially selected for adipose tissue biopsy specimen collection based on FSIGTT results (a power analysis performed before initiation of the study suggested that the number of animals used in this study was appropriate for the measured variables). All tests were performed in April and June 2008. Testing was performed as reported previously by Pratt et al. Briefly, IV catheters were placed in the right and left external jugular veins of each horse after aseptic preparation and subcutaneous (SC) infiltration of the sites with local anesthetic. One catheter was designated for medication administration, and one was designated for blood collection. The horses were confined in stalls overnight after catheter placement to minimize the effect of stress on test results. Blood initially was collected for baseline measurement of plasma insulin and glucose concentrations, followed by administration of a dose of 50% dextrose (300 mg/kg IV, approximately 300 mL, administered over 2–3 minutes). Completion of this infusion marked time ( t ) = 0. Blood samples (approximately 6–8 mL/time point) were collected in EDTA for measurement of insulin and glucose concentrations at 0, 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 12, 14, 16, and 19 minutes postdextrose infusion. A dose of regular insulin (0.1 IU/kg IV) was administered at t = 20 minutes, and additional blood samples were collected at 22, 24, 26, 30, 35, 40, 50, 60, 70, 80, 90, 100, 120, 150, and 180 minutes postdextrose infusion. After completion of sampling, the catheters were removed and the animals returned to the herd. Insulin concentrations were determined by a commercially available radioimmunoassay validated previously for horses, and glucose concentrations were analyzed with a colorimetric assay. Insulin and glucose data were analyzed by minimal model parameters, and the calculated insulin sensitivity (SI; L/min/mU) was used to categorize each horse as insulin resistant (IR; SI < 1.0 L/min/mU) or insulin sensitive (IS; SI > 1.5 L/min/mU). Based on the SI results, 12 of the 20 tested mares (6 IR and 6 IS) were selected to undergo adipose tissue biopsy specimen collection. Adipose Tissue Biopsy Specimen Collection All biopsy specimen collections were performed in July 2008. Horses were transported to the OSU Veterinary Teaching Hospital, weighed upon arrival, and housed individually in box stalls with access to timothy hay, water, and a trace mineral block ad libitum. A physical examination was performed on each horse upon arrival, and morphometric parameters were recorded (neck circumference, girth circumference, ultrasonographic retroperitoneal fat thickness at the umbilicus). All animals were considered healthy based on results of physical examination, CBC, and serum biochemical analysis performed on the day of arrival. Evidence of chronic pyometra, however, was detected in 1 IR mare during hospitalization, and data from this mare therefore were excluded from the final analysis. An acclimation period of 48 hours was observed before biopsy specimen collection. On the day of biopsy specimen collection, a 14 G IV catheter was aseptically inserted in the left external jugular vein for the purposes of sample collection and induction and maintenance of general anesthesia. A pulmonary artery catheter was aseptically inserted via the right external jugular vein for use in collecting blood gas data for a separate study. A tetanus toxoid booster was administered in the left cervical muscle before biopsy specimen collection, and sulfamethoxazole‐trimethoprim (15 mg/kg PO q12h) and phenylbutazone (1 g PO q12h) were administered to each animal (continued for 5 days after biopsy specimen collection). After sedation with xylazine (0.5 mg/kg IV), general anesthesia was induced with ketamine (2.2 mg/kg IV) and diazepam (0.1 mg/kg IV) and maintained with guiafenesin ‐based total IV anesthesia (guiafenesin 5%; xylazine 0.05%; ketamine 0.2%; 1–2 mL/kg/h). All mares were orotracheally intubated, and positive‐pressure ventilation was initiated and maintained with 100% oxygen for the duration of the procedure (approximately 30 minutes). Incisional biopsy specimens of nuchal crest and tail head adipose tissue were collected (∼3 g tissue) with the animal in lateral recumbency after aseptic preparation of the biopsy specimens sites with chlorhexidine surgical scrub and isopropyl alcohol ; incisions were closed in a single layer with 0 polydioxanone monofilament suture material in a simple continuous pattern. Each horse then was placed on a padded surgical table in dorsal recumbency and moved into a surgical suite; the abdomen was aseptically prepared and draped for ventral midline celiotomy. A small ventral midline abdominal incision was created (∼8 cm in length), and adipose tissue biopsy specimens (∼3 g each) were collected from the margins of the incision (retroperitoneal adipose tissue), the mesocolon of the descending colon (mesenteric adipose tissue), and the omentum. The abdominal incision was closed in 2 layers: the linea alba was apposed with 2 polyglactin 910 in a simple continuous pattern, and the skin was apposed with 0 polydioxanone in a simple continuous pattern. Horses then were allowed to recover from general anesthesia and returned to box stalls. All mares remained hospitalized for 10 days after biopsy specimen collection, during which time they were continuously monitored for incisional and other complications. After this period, all mares were returned to the teaching herd. Sutures were removed manually from the nuchal ligament and tail head incisions 14 days after biopsy specimen collection; sutures were not removed from the abdominal incisions. A portion of each adipose tissue biopsy specimen was flash‐frozen in liquid nitrogen at the time of collection and stored at −80°C for later analysis by real‐time quantitative polymerase chain reaction (RT‐qPCR); the remainder was fixed in neutral buffered formalin and paraffin embedded. RNA Isolation, cDNA Synthesis, and RT‐qPCR Homogenates were made from each adipose tissue sample (1.0–1.5 g) with a tissue disruptor, and total RNA was extracted by a modified Trizol method. cDNA was synthesized from total RNA from each sample (after DNAse treatment ) via reverse transcription (Retroscript ) and stored at −20°C until used for RT‐qPCR analysis. A thermocycler (Roche 480 ) was used to perform RT‐qPCR. Amplification was quantified against external standards using fluorescent format for SYBR Green I as described previously. Primers for TNF‐α, IL‐1β, IL‐6, plasminogen activator inhibitor‐1 (PAI‐1), monocyte chemoattractant protein‐1 (MCP‐1), and the housekeeping genes (β‐actin and β 2 ‐microglobulin) were designed against equine‐specific gene sequences (exon sequences) by computer programs as described previously and were used for amplification of the respective genes of interest. Standard curves for quantification of transcript in adipose tissue samples were generated with serial dilutions of a linearized vector containing an insert of each amplified cDNA fragment. Data Analysis RT‐qPCR data from housekeeping genes (β‐actin and β 2 ‐microglobulin) were entered into a computer program to test each gene's quality as a housekeeping gene for equine adipose tissue. Because both genes were determined to be satisfactory by the program, a geometric mean was obtained from the 2 genes' data in order to generate a normalization factor for gene expression from each adipose tissue sample. Gene expression data from the genes of interest then were normalized using this factor for each individual tissue. Data analysis was performed by a statistical software program. Morphometric and gene expression data were assessed for normality by the Shapiro‐Wilk and D'Agostino and Pearson's omnibus normality tests. Morphometric data were analyzed with a Student's t ‐test; gene expression data (or log transforms of nonnormal data) were subjected to 2‐way analysis of variance with repetition on 1 factor (depot) followed by Bonferroni's posttest. Correlation analyses (Pearson's rank‐correlation test) were performed to identify relationships between morphometric parameters and nuchal ligament cytokine gene expression. Statistical significance was accepted at P < .05. Results Differences in age, body weight, body condition score, neck circumference, girth circumference, retroperitoneal adipose tissue thickness, or basal insulin concentration between the IR and IS groups were not detected ( Table 1 ). Subjectively, the omental adipose tissue depot of all horses appeared grossly small at laparotomy. Of the parameters evaluated, the only statistically significant difference between the groups was peripheral SI (IR group, SI = 0.58 ± 0.31 × 10 −4 L/min/mU, n = 5; IS group, SI = 2.59 ± 1.21 × 10 −4 L/min/mU, n = 6; P = .008). No statistically significant difference was observed in acute insulin response to glucose (AIRg) between the groups (IR, AIRg = 821.8 ± 585.8 mU/L/min; IS, AIRg = 394.2 ± 162.4 mU/L/min; P = .18). No correlation was observed between basal insulin concentration and SI ( P = .28). 1 Morphometric characteristics of insulin‐resistant (IR) and insulin‐sensitive (IS) light breed mares from which adipose tissue biopsies were collected (mean ± SD). Group Age (years) Body Weight (lb) Body Condition Score (1–9) Neck Circumference (cm) Girth Circumference (cm) Retroperitoneal Fat Thickness (cm) Basal Insulin Concentration (mIU/L) IR (n = 5) 16.8 ± 4.5 1,122 ± 56 7.2 ± 0.7 90.1 ± 4.2 186.3 ± 4.3 3.0 ± 0.9 22.9 ± 9.9 IS (n = 6) 10.8 ± 7.2 1,193 ± 133 6.75 ± 1.6 93.2 ± 3.1 189.1 ± 6.4 2.7 ± 1.7 15.5 ± 11.9 a Thickness of the retroperitoneal adipose tissue measured ultrasonographically at the level of the umbilicus with a 7.5 mHz linear array probe. The average of 5 measurements was recorded for each horse. No statistically significant effect of SI status on concentrations of mRNA coding for TNF‐α ( P = .29), IL‐1β ( P = .80), IL‐6 ( P = .93), MCP‐1 ( P = .23), or PAI‐1 ( P = .72) was noted within each adipose tissue depot ( Fig 1 ). However, a significant difference in gene expression among depots was observed, with higher concentrations of IL‐6 ( P = .004) and IL‐1β ( P = .0005) mRNA measured in nuchal ligament adipose tissue compared with other depots ( Fig 2 ). No differences in gene expression of TNF‐α ( P = .97), MCP‐1 ( P = .14), or PAI‐1 ( P = .07) were observed among depots ( Fig 2 ). 1 Gene expression of IL‐1β, IL‐6, TNF‐α, MCP‐1, and PAI‐1 by adipose tissue depot. Data from IR and IS groups have been stratified for each depot. No statistically significant differences in gene expression were observed between IR and IS groups for each depot. Prominent individual variability in gene expression was noted in IR and IS animals. OM, omental; RET, retroperitoneal; NUC, nuchal ligament; MES, mesocolonic; TAIL, tail head; TNF‐α, tumor necrosis factor‐α; IL, interleukin; PAI‐1, plasminogen activator inhibitor‐1; MCP‐1, monocyte chemoattractant protein‐1; IR, insulin resistant; IS, insulin sensitive. 2 Gene expression of IL‐1β, IL‐6, TNF‐α, MCP‐1, and PAI‐1 by adipose tissue depot. Data from IR and IS groups have been pooled for each depot. Expression of IL‐1β and IL‐6 is significantly higher in the nuchal ligament adipose tissue depot (*); no other statistically significant differences were observed. OMEN, omental; RETRO, retroperitoneal; NUCHAL, nuchal ligament; MESEN, mesocolonic; TAIL, tail head; TNF‐α, tumor necrosis factor‐α; IL, interleukin; PAI‐1, plasminogen activator inhibitor‐1; MCP‐1, monocyte chemoattractant protein‐1; IR, insulin resistant; IS, insulin sensitive. There was a significant correlation between the expression of IL‐1β and IL‐6 in the nuchal ligament ( r = 0.76; P = .006). However, there was no statistically significant correlation between IL‐1β or IL‐6 expression and mean neck circumference (IL‐1β: r = 0.24, P = .48; IL‐6: r = 0.31, P = .36), body condition score (IL‐1β: r =− 0.10, P = .76; IL‐6: r = 0.11, P = .74), and body weight (IL‐1β: r = 0.08, P = .82; IL‐6: r = 0.15, P = .64). Discussion The study described here compares proinflammatory gene expression profiles of multiple visceral and SC adipose tissue depots of light breed horses. The results suggest that the visceral adipose tissue depots of the adult horse do not have greater expression of genes encoding inflammatory cytokines when compared with SC adipose tissue depots, a finding that differs substantially from observations made in other species. Omental adipose tissue, which has been reported to be the primary depot undergoing inflammatory signaling in obese humans and laboratory animals, and where fat accumulation reportedly correlates closely with cardiovascular disease risk in obese humans, does not appear to display a greater inflammatory gene expression profile than other adipose tissue depots in the adult horse. EMS has emerged in recent years as a substantial risk factor for development of endocrinopathic laminitis in horses and ponies. Whereas systemic insulin resistance, increased adiposity (particularly regional adiposity), hyperinsulinemia, and dyslipidemia characterize the syndrome as it is currently accepted, only insulin resistance has thus far been identified as an independent risk factor for laminitis and directly involved in its pathogenesis. The horses used in this study displayed no physical evidence of prior bouts of laminitis; neither did they have prominent nuchal ligament adipose tissue deposits. These findings suggest that these subjects should not be categorized as having EMS per se. However, because insulin resistance may be the component of EMS most directly associated with laminitis risk, its relationship to proinflammatory cytokine and chemokine gene expression within the adipose tissue was investigated. In humans and experimental rodent models of obesity, adipose tissue (particularly visceral depots) expresses higher concentrations of TNF‐α, IL‐1β, IL‐6, and other proinflammatory cytokines and chemokines in peripherally IR individuals than in those that are IS. Interestingly, the data from the present study do not identify any differences in adipose tissue inflammatory gene expression between depots from IR and those from IS horses of similar body condition scores (all animals being overweight). However, differences in IL‐1β and IL‐6 gene expression among adipose tissue depots were statistically significant, suggesting that the nuchal crest is a more active depot. The lack of an observed effect of peripheral SI on proinflammatory gene expression within a specific depot suggests that the differences noted among depots may be associated with other factors related to obesity. An important confounder in the interpretation of these results is the fact that all of the horses sampled were either overweight or obese according to a widely used body condition scoring system. In several studies of horses, investigators have demonstrated positive correlations between body condition score and peripheral insulin resistance, plasma basal insulin concentration, plasma leptin concentration, proinflammatory cytokine gene expression, and protein concentration in peripheral blood, and risk of laminitis. Negative correlations have been suggested between body condition score and plasma adiponectin concentration in horses, a finding in keeping with this adipokine's reported anti‐inflammatory and cardioprotective value in the setting of human metabolic syndrome. In humans, it is well recognized that obese individuals often are IR. In fact, body mass index and insulin resistance each have been interpreted as proxies for the other in certain assessments of cardiometabolic risk. However, the existence of peripherally IS obese individuals is difficult to explain in this setting. McLaughlin and colleagues recently reported that although, predictably, obesity was more prevalent in a cohort of IR individuals (34%) than lean body composition (16%), obesity and insulin resistance did not make equivalent contributions to combined cardiovascular disease risk in their risk‐assessment model. Their data reinforce the concept that although body mass index and insulin resistance are related, they are not synonymous and contribute independently to cardiovascular risk. Obesity and insulin resistance clearly are correlated in humans and horses, but their respective contributions to subsequent pathology cannot be assumed to be equal. Furthermore, the recent description of a “metabolically healthy but obese” phenotype in several human study populations, in which obesity is notably not accompanied by insulin resistance, hyperglycemia, dyslipidemia, or an increased risk of cardiovascular disease, suggests uncoupling of obesity and metabolic risk in a substantial proportion of individuals. For example, a recent large survey of humans identified 31.7% of obese adults as “metabolically healthy” and 23.5% of lean adults as “metabolically abnormal.” Additionally, very recent work in mice identifies a candidate gene, Brd2, as a molecular link between obesity and insulin resistance. Brd2 knockout mice develop severe obesity and hyperinsulinemia, but they display enhanced glucose tolerance, decreased adipose tissue macrophage infiltration, and increased plasma concentrations of adiponectin. Furthermore, these mice displayed evidence of enhanced expression of proinflammatory cytokines that appeared unrelated to their systemic SI or the degree of adipose tissue macrophage infiltration. Thus, examples from multiple species have weakened the assertion that obesity and insulin resistance are inexorably related. This information may in part explain why a relationship between insulin resistance and gene expression was not identified in this study and why evidence of increased nuchal ligament inflammatory mediator expression was noted in all mares. Therefore, adiposity may factor more importantly in EMS and endocrinopathic laminitis risk than solely through its association with insulin resistance. To further test this possibility, lean cohorts of animals should be included in similar studies in the future to assess whether inflammatory signaling occurs with obesity in the horse regardless of SI. University‐owned mares were an attractive source population for the study reported here because of their ready availability and standardized, uniform management (feeding, housing, and preventive medicine). However, sex influences on both peripheral SI and biological behavior of the adipose tissue have been noted in horses and humans. Additionally, data exist to support the contention that mares have an increased risk of (all‐cause) laminitis than do stallions or geldings, implicating an influence of sex on the most concerning complication of equine obesity. We recognize this influence as a confounder to the interpretation of our results and acknowledge that the data reported here may not be representative of what might be noted in a population of horses including both sexes (intact and altered). An important finding of this study, that nuchal crest fat appears to be the most reactive fat depot (with respect to inflammatory signaling) in the horse, complements clinical findings of previous investigators who have identified the accumulation of nuchal crest adipose tissue as a risk factor for laminitis associated with equine obesity (global, regional, or both). Frank et al reported that obese horses with insulin resistance had greater mean neck circumference scores than nonobese mares, relating this morphologic change with the SI status and body condition of the horse. Accumulation of nuchal ligament adipose tissue has further been shown to have predictive value in assessing risk of pasture‐associated laminitis in multiple cohorts of ponies, with ponies that have larger neck crests being at greater risk for developing the condition. Thus, our finding of unique biological behavior of nuchal crest fat when compared with other depots supports an accumulating body of evidence that the morphology of the nuchal crest changes in the setting of EMS and is correlated with clinical signs of the disease. In conclusion, the current study does not support a direct relationship between the degree of insulin resistance and inflammatory mediator gene expression by adipose tissue in chronically obese horses, suggesting that other factors besides insulin resistance may mark risk for laminar injury in the obese horse. Additionally, the results of the current study indicate that the nuchal ligament adipose depot, rather than visceral adipose depots, may be more important pathophysiologically in the horse. This finding not only fits well with the clinical case at risk of laminitis with enlarged nuchal adipose deposits (ie, a “cresty neck”), but also indicates that it may be possible to focus future adipobiology studies in horses on nuchal ligament tissue biopsy specimens (versus obtaining abdominal or tail head samples). Additional study will be required to determine the role of the adipose tissue in EMS, including the link between changes in the adipose tissue and laminitis and its potential as a therapeutic target in animals at risk of EMS‐associated laminitis. Footnotes a Abbocath 14 G, Abbott Medical Health, Abbott Park, IL b Dextrose 50%, Vedco, St Joseph, MO c BD Vacutainer K 2 EDTA tubes, BD (Becton Dickinson), Franklin Lakes, NJ d Humulin R, Eli Lily and Company, Indianapolis, IN e Coat‐A‐Count insulin RIA, Siemens Medical Solutions Diagnostics, Los Angeles, CA f Thermo‐Trace Ltd, Melbourne, Australia g Tetanus toxoid, Fort Dodge Animal Health/Wyeth, Madison, NJ h Sulfamethoxazole‐trimethoprim 960 mg tablets, Mutual Pharmaceutical Co Inc, Philadelphia, PA i Phenylzone Paste, Schering Plough Animal Health, Union, NJ j Xylazine HCl, 100 mg/mL, IVX Animal Health Inc, St Joseph, MO k Ketaset, Fort Dodge Animal Health/Wyeth l Diazepam Inj, 5 mg/mL, Abbott Laboratories m Guifenesin 5%, Butler Animal Health Supply, Dublin, OH n Chlorhexidine 2% surgical scrub, First Priority Inc, Elgin, IL o Isopropyl alcohol 70%, Butler Animal Health Supply p PDS, Ethicon Inc, Somerville, NJ q Vicryl, Ethicon Inc r Tissue‐Ruptor, Qiagen, Valencia, CA s DNAse I, Thermo Fisher Scientific Inc, Rockford, IL t Retroscript, Ambion Inc, Austin, TX u Roche 480, Roche Applied Science, Indianapolis, IN v TOPO TA Cloning Kit, Invitrogen, Carlsbad, CA w geNorm, Ghent University, Ghent, Belgium x GraphPad Prism 5, GraphPad Software, La Jolla, CA Acknowledgments This work was supported by a grant from the American Quarter Horse Association. The authors acknowledge the following individuals for their assistance with this project: Dr Nicholas Frank and Sarah Elliot (University of Tennessee) for provision of the RNA extraction protocol; Drs Turi Aarnes and Yukie Ueyama for anesthesia support; Drs Britta Leise, Rafael Faleiros, Jarred Williams, Ms Cailing Yin, and Drs Kelly Welin and Trina Westerman for assistance with biopsy specimen collection; and Ms Mauria Watts for critical review of the manuscript.

Journal

Journal of Veterinary Internal MedicineWiley

Published: Jul 1, 2010

Keywords: ; ; ;

References

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