Obesity has reached epidemic proportions in humans and companion animals, including horses.1–3 Obesity is a risk factor for higher morbidity and mortality rates in humans,3 laboratory animals with experimentally induced obesity,4 and horses.5,6 Furthermore, obesity appears to compound the inflammatory and cytokine responses to acute conditions (eg, sepsis) and may lead to a state of chronic inflammation.6
Obesity has been associated with greater permeability of gastrointestinal mucosa in humans7–9 and mice.10–13 Altered permeability may allow absorption of bacterial toxins as well as translocation of bacteria, which can result in local inflammation and life-threatening sequelae such as activation of the systemic inflammatory response and disseminated sepsis. Integrity of the gastrointestinal mucosa is of particular concern in horses; they have vast amounts of bacteria in the large intestine for fermentation of forage, and they are exquisitely sensitive to LPS.14 A relationship between obesity and chronic elevations in concentrations of proinflammatory cytokines has been established in humans,15–17 and a similar association has been suggested in horses.18–20 In obese people, adipocyte secretion of tumor necrosis factor-α attracts macrophages to adipose tissue. Once activated, these macrophages release additional proinflammatory cytokines.21–23 Proinflammatory cytokines may contribute to dysfunction of the intestinal barrier by affecting the structure and localization of mucosal tight junction proteins, specifically zonula occludens-1 and occludin.13
Furthermore, diet and endocrine status may have additional effects on gastrointestinal permeability in obese animals. For example, obese mice fed a high-fat diet had higher concentrations of circulating LPS, compared with the concentrations in obese mice fed a lower-fat diet, and the high-fat diet was associated with development of insulin dysregulation.24
The primary objective of the study reported here was to determine whether obese horses would have greater permeability of the intestinal mucosa, compared with that of lean horses. Specifically, we hypothesized that obese horses would have a lower mucosal TER and greater transmucosal flux of LPS, both of which are consistent with greater mucosal permeability, when compared with results for lean horses. A secondary objective was to assess whether alterations in mucosal permeability would be associated with insulin dysregulation.
Materials and Methods
Animals
Thirteen client-owned horses that were to be euthanized for reasons unrelated to the gastrointestinal tract were recruited for the study. There were 7 obese horses (BCS, 7 to 9 [scale, 1 to 9]) and 6 lean horses (BCS, 4 or 5). Horses ranged from 8 to 15 years of age and had no history of gastrointestinal tract disease or treatment with NSAIDs or antimicrobial medications within the preceding 30 days. Owners provided informed consent for donation of their horses for the study. All procedures were approved by the Michigan State University Animal Care and Use Committee.
Each horse was weighed and the BCS assessed. The BCS of each horse was assigned separately by each of 3 trained investigators by use of a method described elsewhere.25 Horses were then housed in a lot with a dirt surface for approximately 14 days. They had unlimited access to mixed grass hay and water, but no concentrate feeds were provided.
Metabolic testing
After a minimum of 5 days during the 14-day acclimation period, an OST was performed on each horse to characterize glucose and insulin dynamics.26 Briefly, horses were housed in a stall overnight, and food was withheld for 8 hours. Corn syrupa was administered (15 mL/100 kg, PO). Blood samples (10 mL/sample) were collected via jugular venipuncture with a 20-gauge, 1.5-inch needle into serum and heparinized evacuated tubes immediately before (time 0) and 15, 30, 45, 60, 75, 90, 105, 120, 150, and 180 minutes after administration of corn syrup. Heparinized blood samples were placed on ice and centrifuged within 30 minutes after collection, whereas serum tubes were allowed to clot at ambient temperature and then were centrifuged within 2 hours after collection. Supernatant plasma and serum were transferred to plastic tubes and frozen at −20°C until batch analysis of glucose and insulin concentrations.
Glucose concentrations were measured in duplicate by use of a membrane-based glucose oxidase method.b Insulin concentrations were measured in duplicate by use of a radioimmunoassayc as described elsewhere.27 Intra-assay and interassay coefficients of variation for serum with low and high insulin concentrations ranged between 1.7% and 11.8%. Horses were classified as having insulin dysregulation when the serum insulin concentration exceeded 45 mU/mL from 60 to 90 minutes after administration of corn syrup.28
Collection of tissue samples
After the 14-day acclimation period was completed, horses were weighed again to ensure that body weight had not changed by ≥ 2%. A catheter was placed in a jugular vein, and horses were euthanized with sodium pentobarbitald (95 mg/kg, IV). Death was confirmed, and an incision was made on the ventral midline. Full-thickness tissue samples were obtained from the midjejunum, distal portion of the ileum, midcecum (30 cm from the apex of the cecum), right dorsal colon, and pelvic flexure. Mucosal samples were obtained from the rectum. All samples were immediately placed in oxygenated lactated Ringer solution and maintained at 37°C.
Measurement of TER
Within 15 minutes after each horse was euthanized, the mucosa for each intestinal sample was carefully dissected from the seromuscular layer and mounted in duplicate Ussing chambers.e The TER was measured as described elsewhere.29 Briefly, tissues were bathed on the luminal and serosal sides with 10 mL of oxygenated lactated Ringer solution. In addition, the solution bathing the serosal side contained glucosef (10 mmol/L), and the solution bathing the luminal side was osmotically balanced by the addition of mannitolf (10 mmol/L). Bathing solutions were circulated and maintained at 37°C by use of water-jacketed reservoirs. Tissues were allowed a 15-minute equilibration period in the Ussing chamber. Potential difference then was measured by use of Ringer-agar bridges connected to calomel electrodes, and the short-circuit current was measured by use of an automated voltage clamp. Electrical measurements were recorded every 15 minutes for 3 hours. The TER was calculated with the short-circuit current and potential difference by use of Ohm's law.
Measurement of LPS flux
After a 30-minute equilibrium period was completed, 83 μg of FITC-labeled LPSf (Escherichia coli O55:B55) was added to the solution bathing the luminal side of each intestinal segment, in accordance with a method described elsewhere.30 Ussing chambers used for these evaluations were protected from light. Samples (200 μL) of bathing solution were obtained in triplicate from both the luminal and serosal sides immediately before and 120 minutes after addition of FITC-LPS. Dilutions of FITC-LPS (0 to 5 ng/mL) were used to construct a standard concentration curve, and fluorometry was used to determine the concentration of FITC-LPS in the samples. The LPS flux to the serosal side of the tissue was calculated.
Histologic examination
Full-thickness biopsy samples from the jejunum, ileum, cecum, pelvic flexure, and right dorsal colon and rectal mucosa samples, all of which were obtained at locations adjacent to the sites of the samples obtained for the Ussing chambers, were examined by use of light microscopy. Three 5-μm tissue sections for each intestinal segment were aligned on the villus-crypt axis and stained with H&E stain. Height and width of 3 jejunal and ileal villi and depth of 3 crypts in each segment were measured with a micrometer in the light microscope; measurements were made 3 times by 1 investigator who was unaware of the tissue source. Total villus surface area was estimated on the basis of the formula used to calculate surface area of a cylinder as follows: villus surface area = 2π• (0.5•[{4/π}/w])•h, where w is villus width and h is villus height. Interstitial-to-crypt ratios for the cecum, pelvic flexure, right dorsal colon, and rectum were calculated by determining the mean interstitial distance of the three 5-μm tissue sections and then dividing this value by the mean value of the crypt width for each intestinal segment, as described elsewhere.31
Statistical analysis
Horses were grouped on the basis of BCS as obese or lean. Values for BCS, glucose and insulin concentrations, and LPS flux and histologic data were tested for normality by use of Shapiro-Wilk analyses. Normally distributed data (BCS, LPS flux, and histologic data for each intestinal segment) were compared between groups by use of a t test, whereas non-normally distributed data were compared by use of a Mann-Whitney U test. Data for TER were assessed by use of a 2-factor repeated-measures ANOVA, with BCS group and intestinal segment as the factors. All analyses were performed with statistical software.g Significance was set at values of P < 0.05.
Results
BCS and OST
The BCS was significantly (P = 0.01) greater for obese (median, 8.0; range, 7 to 9) than lean (median, 4.25; range, 4 to 5) horses. Dynamic glucose and insulin testing revealed results supportive of insulin dysregulation for 5 of 7 obese horses and 1 of 6 lean horses (Table 1). Results for the OST were not available for 1 obese horse.
Insulin concentrations of lean and obese horses determined by use of an OST.
| Insulin concentration (μU/mL) | |||||||
|---|---|---|---|---|---|---|---|
| 60 minutes | 90 minutes | ||||||
| Group | No. of horses | Median | SEM | Range | Median | SEM | Range |
| Lean | 6 | 40.7 | 4.5 | 29.5–59.7 | 37.8 | 6.0 | 25.5–71.5 |
| Obese | 6* | 67.4 | 12.6 | 18.1–116.5 | 67.4 | 12.7 | 25.1–109.5 |
| Without insulin dysregulation | 6† | 33.9 | 3.8 | 18.1–43.8 | 33.6 | 2.78 | 25.5–43.2 |
| With insulin dysregulation | 6† | 67.4 | 9.6 | 38.5–116.5 | 79.1 | 9.8 | 48.0–109.5 |
Lean horses had a BCS of 4 or 5 (scale, 1 to 9), and obese horses had a BCS of 7 to 9. Horses were classified as having insulin dysregulation when the serum insulin concentration exceeded 45 mU/mL from 60 to 90 minutes after oral administration of corn syrup.
Represents results for only 6 horses; insulin concentrations were not available for 1 obese horse.
There was 1 horse in the obese group without insulin dysregulation and 1 horse in the lean group with insulin dysregulation.
TER
The TER did not differ significantly between obese and lean horses in the jejunum, ileum, cecum, pelvic flexure, right dorsal colon, and rectum (Figure 1). Similarly, when horses were classified on the basis of insulin dysregulation status, no differences in TER were found in any segments of the intestinal tract between horses with or without insulin dysregulation (data not shown).
Mean ± SEM values of TER for sections of mucosa obtained from the jejunum (A), ileum (B), cecum (C), pelvic flexure (D right dorsal colon (E), and rectum (F) of 7 obese horses (BCS, 7 to 9 [scale, 1 to 9]; dashed line) and 6 lean horses (BCS, 4 to 5; sol line). Notice that the y-axis scale in panel C differs from the y-axis scale of the other panels. Values did not differ significantly (P ≥ 0.0 between obese and lean horses in any intestinal segment.
Citation: American Journal of Veterinary Research 80, 8; 10.2460/ajvr.80.8.792
Mean ± SEM values of TER for sections of mucosa obtained from the jejunum (A), ileum (B), cecum (C), pelvic flexure (D right dorsal colon (E), and rectum (F) of 7 obese horses (BCS, 7 to 9 [scale, 1 to 9]; dashed line) and 6 lean horses (BCS, 4 to 5; sol line). Notice that the y-axis scale in panel C differs from the y-axis scale of the other panels. Values did not differ significantly (P ≥ 0.0 between obese and lean horses in any intestinal segment.
Citation: American Journal of Veterinary Research 80, 8; 10.2460/ajvr.80.8.792
Mean ± SEM values of TER for sections of mucosa obtained from the jejunum (A), ileum (B), cecum (C), pelvic flexure (D right dorsal colon (E), and rectum (F) of 7 obese horses (BCS, 7 to 9 [scale, 1 to 9]; dashed line) and 6 lean horses (BCS, 4 to 5; sol line). Notice that the y-axis scale in panel C differs from the y-axis scale of the other panels. Values did not differ significantly (P ≥ 0.0 between obese and lean horses in any intestinal segment.
Citation: American Journal of Veterinary Research 80, 8; 10.2460/ajvr.80.8.792
LPS flux
After samples were incubated for 120 minutes, LPS flux across jejunal mucosa collected from obese horses was significantly (P = 0.016) greater than that for jejunal mucosa collected from lean horses (Table 2). There were no significant differences in LPS flux across the mucosa of the ileum, cecum, right dorsal colon, and rectum between obese and lean horses. Data for LPS flux were not available for the pelvic flexure mucosa because of technical difficulties. No significant differences in LPS flux were found when comparing horses with and without insulin dysregulation (data not shown).
Mean ± SEM values for total LPS flux (ng of LPS/h/cm2) across intestinal mucosa obtained from lean and obese horses.
| Intestinal segment | No. of horses* | Lean | Obese | P value† |
|---|---|---|---|---|
| Jejunum | 6 and 7 | 2.9 ± 1.2 | 17.0 ± 4.6 | 0.016 |
| Ileum | 4 and 7 | 2.1 ± 0.2 | 2.2 ± 0.2 | 0.892 |
| Cecum | 5 and 7 | 3.1 ± 0.5 | 1.4 ± 0.5 | 0.142 |
| Right dorsal colon | 6 and 7 | 9.5 ± 6.2 | 14.4 ± 10.8 | 0.718 |
| Rectum | 6 and 7 | 0.5 (−0.65 to 1.8)‡ | 1.5 (0.5 to 4.2)‡ | 0.181 |
Represents the number of lean and obese horses for which intestinal segments were evaluated.
Values were considered to differ significantly at P < 0.05.
Values reported are median (interquartile [25th to 75th percentile] range).
Histologic examination
No significant differences in villus height, villus width, and surface area of the jejunum and ileum were detected between obese and lean horses. No significant differences were found when comparing interstitial-to-crypt ratios between obese and lean horses for the cecum, pelvic flexure, right dorsal colon, and rectum.
Discussion
In the study reported here, obese horses had greater LPS flux across jejunal mucosa than did lean horses, which indicated an increase in mucosal permeability to LPS. Therefore, our hypothesis was supported. However, when data were reanalyzed to compare horses with and without insulin dysregulation, there were no significant differences.
An association between obesity and greater permeability of the gastrointestinal mucosa, particularly of the small intestine, has been identified in humans7–9 and laboratory animals.10–13 This is consistent with results of the present study in which there was greater LPS flux across the jejunal mucosa in obese horses. There are 2 pathways for transport across the mucosa: the transcellular pathway and the paracellular pathway. Several mechanisms for passage of solutes exist within the paracellular pathway, including a high-capacity, charge-selective pore mechanism that allows for passage of small ions and uncharged molecules and a lower-capacity leak mechanism that allows for flux of larger molecules.32 The paracellular pathway reportedly accounts for 75% to 94% of TER33–35; however, despite a greater LPS flux, there was not a lower TER of jejunal mucosa found in tissues from obese horses, compared with that for tissues from lean horses in present study. One possible explanation for this apparent discrepancy was that LPS flux occurred solely through the low-capacity leak mechanism, and changes in specific interepithelial tight junction proteins may have allowed an increase in LPS flux without a concomitant decrease in TER.32,36,37 Tight junction proteins, including claudins, occludins, and junctional adhesion molecules, are responsible for the control of paracellular transport through both restrictive and nonrestrictive pores.38,39 In addition, concentrations of inflammatory cytokines that are increased with obesity could differentially affect paracellular permeability to specific molecules.13,40 For example, if tight junction proteins responsible for limiting paracellular transport of LPS through restrictive pores are affected to a greater degree by obesity, LPS flux could increase without a substantial impact on TER.41 Investigators of 1 study41 determined that interferon-γ increases permeability of gastrointestinal mucosa to LPS through LPS-specific pores (restrictive) but does not increase permeability of other small molecules. Obesity is associated with an increase in the production of adipose tissue–derived proinflammatory cytokines, which could explain why differences were detected in LPS flux but not TER in the study reported here. The reason that evidence of changes in mucosal permeability was limited to jejunal tissue in the present study and other studies8,11–13 remains unclear.
Three additional mechanisms have also been proposed to explain the association between greater gastrointestinal permeability and obesity: alterations in gastrointestinal microbiota leading to changes in tight junction integrity; dietary effects, particularly for high-fructose and high-fat diets; and nutrient deficiencies.4 The changes in microbial composition of the gastrointestinal tract that may lead to increased permeability in humans and laboratory animals remain unclear. The gastrointestinal microbiome was not evaluated in the horses of the present study. The association between BCS and fecal microbiota in horses has been evaluated in several studies.42–45 Results have been inconsistent, but differences in diversity, but not the presence or absence of specific microorganisms, in the microbiome have been identified.42–44 Although fecal samples are easy to obtain for microbial analysis, they are not likely to be representative of the small intestinal microbiome,46 which was the intestinal location at which we detected differences in permeability in the study reported here. To the authors’ knowledge, no studies have been conducted to evaluate differences in the microbiome of the small intestine between obese and lean horses. In addition, all horses of the present study were housed in dirt lots and fed grass hay for 2 weeks; thus, they were receiving a low-starch, low-fat diet. Finally, although there are many vitamin and mineral products available for horses, the authors are not aware of any specific nutrient deficiency that affects mucosal permeability in horses.1–3
The cause of obesity is complex and includes genetics, diet, and environment.47 In humans, obesity has been associated with an increase in the risk for > 20 medical conditions and is a consistent predictor of higher morbidity and mortality rates.48,49 A similar higher risk of comorbidities has also been found in veterinary species, including dogs,2,50,51 cats,2,50,51 and horses.52 Obesity in dogs is associated with the development of diabetes mellitus,51 hepatic lipidosis,51 early-onset osteoarthritis of the hip joints,53 airway dysfunction (including decreased expiratory function54 and reactive airway disease55), and incompetence of the urethral sphincter mechanism.56 In horses, obesity has been associated with development of laminitis.57 In another study of horses,52 increased expression of adiponectin mRNA in retroperitoneal fat, but not body mass index, was associated with increased risk of death after colic surgery. The association between obesity and morbidity and mortality rates may be attributable, in part, to a link between obesity and insulin dysregulation.58 In laboratory mice, increased gastrointestinal permeability has been associated with higher circulating concentrations of LPS and development of insulin dysregulation.24 The binding of LPS to CD14 receptors appears to play an important role in development of insulin dysregulation in mice because CD14 knockout mice with high circulating concentrations of LPS do not have abnormalities in insulin sensitivity and dynamics.24 In general, obesity in horses is frequently associated with insulin dysregulation.59 Five of 7 horses classified as obese and 1 of 6 horses classified as lean in the study reported here had concurrent insulin dysregulation, as determined on the basis of OST results. The interval between the start of the study and performance of the OST was chosen to provide an acclimation period and thus limit the possible effect of stress on OST results. Interestingly, the lean horse that was classified as obese on the basis of OST results had the longest acclimation period; this horse had been housed at our university as part of another unrelated unpublished study. It is interesting that when horses were classified on the basis of insulin dysregulation status without regard to BCS, no significant differences were detected in TER or LPS flux. Clearly, cause-and-effect relationships between obesity, systemic increases in concentrations of proinflammatory cytokines, gastrointestinal permeability, and insulin dysregulation are complex and remain to be fully elucidated.
Limitations of the study reported here included a small number of horses in each group and a diet-acclimation period of only 14 days. Group size for the present study was estimated by use of data from a study13 of mice that was conducted to assess the association between obesity and gastrointestinal permeability; similar data were not available for horses. For α = 0.05, power estimates for TER and LPS flux data were as low as 0.05 and 0.125, respectively, for some intestinal segments. To achieve a desired power of 0.8, larger group sizes would have been needed to avoid type 2 errors. Nevertheless, a difference in LPS flux was detected in jejunal tissue collected from obese and lean horses, which supported the need for further investigation of the potential effects of obesity on gastrointestinal permeability in horses. Changes in diet can induce alterations in the microbiota of humans (which could affect gastrointestinal permeability) in as few as 6 days.60 Because of a more prolonged gastrointestinal transit time in horses, a 14-day diet-acclimation period was chosen for the present study, but this may not have fully eliminated potential confounding effects of variations in diets before study enrollment.60 Although the protocol for recruiting study horses included confirmation by owners that the horses had not received medications during the 30 days before enrollment, prior medication use could have had long-standing effects on gastrointestinal permeability that may also have affected the results of the present study. In addition, measurement of circulating concentrations of inflammatory cytokines, adipokines, and LPS would have better characterized potential effects of obesity in horses but was beyond the scope of this study. Use of the OST to characterize insulin dysregulation in horses also has limitations,61,62 but it was considered the most practical screening test for horses enrolled in the study. Evaluation of specific tight junction proteins, as described in another study,36 may have provided additional mechanistic insight into the changes detected in LPS flux, but not TER, in jejunal mucosa. Assessing flux across the mucosa by use of additional markers, such as 3H-labeled mannitol or 3H-labeled inulin,36,37 may also have allowed us to make conclusions about altered transport via restrictive and nonrestrictive paracellular pores. Finally, we specifically grouped horses as obese or lean by use of the simple measure of BCS, rather than on the basis of estimates of body fat mass (eg, deuterium dilution). This approach was used because BCS is the most clinically relevant assessment of obesity available to veterinarians who may be faced with considering the influence of obesity on medical problems.
To the authors’ knowledge, the study reported here was the first study conducted to assess the effect of obesity on gastrointestinal permeability in horses. We found that obese horses had greater jejunal LPS flux than did lean horses, which would be consistent with greater permeability of the jejunal mucosa via the paracellular route. This finding suggested that obese horses may be at increased risk from greater systemic absorption of LPS. Further studies are warranted to more fully understand alterations in gastrointestinal permeability that could impact morbidity and mortality rates in obese horses.
Acknowledgments
Supported by funds from the Michigan State University College of Veterinary Medicine Endowed Research Funds, specifically the Freeman Fund for Equine Research.
Presented in abstract form at the International Veterinary Emergency and Critical Care Forum, Grapevine, Tex, September 2016.
The authors thank Sue Wismer for technical assistance and care of the horses and Dr. Patty Webber for technical assistance with glucose and insulin testing.
ABBREVIATIONS
| BCS | Body condition score |
| FITC | Fluorescein isothiocyanate |
| LPS | Lipopolysaccharide |
| OST | Oral sugar test |
| TER | Transepithelial resistance |
Footnotes
Karo Syrup Light, ACH Food Co Inc, Oak Brook, Ill.
YSI 2300 STAT plus glucose and lactate analyzer, YSI Inc, Yellow Springs, Ohio.
MP Biomedicals, Santa Ana, Calif.
Vortech Pharmaceuticals Ltd, Dearborn, Mich.
Physiologic Instruments, San Diego, Calif.
Millipore Sigma, St Louis, Mo.
SigmaStat, Systat Software Inc, San Jose, Calif.
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