Numerous stresses such as weaning, infection, and inflammation can lead to intestinal mucosal injury in pigs,1–3 which may result in diarrhea and poor growth of pigs. Therefore, modulation of injured intestines is extremely important in compromised pigs.
Peroxisome proliferator-activated receptors, members of the nuclear hormone receptor superfamily, are ligand-activated transcription factors that include PPARα, PPARβ, and PPARγ4 The PPARs regulate gene expression by binding (with the retinoid X receptor as a heterodimeric partner) to specific DNA sequences, which are termed PPAR response elements.5 Adipose tissue and components of the immune system are the primary sites of PPARγ expression.4,6 In addition, PPARγ is expressed in the small intestines and colon.3,7 Activation of PPARγ is via both natural (ie, 15-deoxy-prostaglandin-J2 and certain polyunsaturated fatty acids) and synthetic (ie, the thiazolidinediones) ligands.4,8 The PPARγ ligands are involved in a number of experimentally induced inflammatory bowel diseases, such as acute ulcerative colitis induced by intracolonic administration of TNBS in rats,9 dextran sodium sulfateinduced colitis in mice,10 and ischemia-induced colitis in rats,7 as well as in ulcerative colitis in humans.11 Although data are available to support a role for PPARγ in the regulation of intestinal inflammation in humans and rodents,8,9,12 few studies have been conducted to investigate these effects in weaned pigs. Investigators in 1 study3 detected PPARγ mRNA expression in the small intestines of weaned pigs. This finding provides a precedence for further study to determine the potential role of PPARγ in weaned pigs with intestinal injury. To examine the potential role of PPARγ in pigs, the study reported here was conducted to investigate whether rosiglitazone, a characterized PPARγ agonist,13,14 could alleviate intestinal injury induced by administration of Escherichia coli LPS in weaned pigs.
Materials and Methods
Animals—Eighteen weaned crossbred (Duroc X Large White X Landrace) pigs (mean ± SD age, 28 ± 3 days; mean body weight, 10.06 ± 0.38 kg) were included in the study. Each pig was housed separately in a 1.80 × 1.10-m2 pen in rooms maintained at a temperature of 25° to 27°C with natural lighting. Pigs were allowed ad libitum access to feed and water. Pigs were fed a conventional diet formulated for weaned pigs during a 14-day adaptation period. Before the study, all pigs had no clinical signs or laboratory evidence of enteric diseases. The animal protocol for this study was approved by the Animal Care and Use Committee of Hubei Province.
Experimental design—Pigs were allocated on the basis of initial body weight and breed to form 3 equivalent treatment groups. Pigs in the control group were injected IP with 10% DMSO (rosiglitazone vehicle) and then 30 minutes later were injected IP with sterile saline (0.9% NaCl) solution. Pigs in the LPS-challenged group were injected IP with 10% DMSO and then 30 minutes later were injected with E coli serotype O55:B5 LPSa (100 μg/kg, IP). Pigs in the rosiglitazone plus LPSchallenged group were injected with rosiglitazoneb (3 mg/kg, IP) and then 30 minutes later were injected with E coli serotype O55:B5 LPS. The LPS was dissolved in sterile saline solution (500 mg of LPS/L of saline solution). The rosiglitazone was dissolved in 10% DMSO (vol/vol). The doses of rosiglitazone and LPS were selected on the basis of the reports of Cuzzocrea et al15 and Liu et al,3 respectively. At 3 hours after challenge exposure, all pigs in the 3 groups were euthanized by IV injection of sodium pentobarbital (40 mg/kg), and samples of intestinal tissues were collected. The time of 3 hours after LPS injection was chosen for tissue collection on the basis of a study16 in which investigators found that there was severe damage to the intestinal mucosa in pigs at 3 hours after LPS administration.
Sample collection—Immediately after each pig was euthanized, the abdominal cavity was opened and the small intestines were removed. Segments (10 cm and 2 × 3 cm) were obtained at points located at 25%, 50%, and 75% of the total intestinal length to represent samples from the duodenum, jejunum, and ileum, respectively. The 10-cm intestinal segments were incised longitudinally, and the contents were flushed with icecold PBS solution (pH, 7.4). The mucosa was scraped with a glass slide, snap-frozen in liquid nitrogen, and stored at –80°C until analysis of disaccharidase activity. The 2 × 3-cm intestinal segments were flushed gently with ice-cold PBS solution and fixed in 4% paraformaldehyde solution for histologic examination.
Intestinal morphology measurements—Tissue specimens were fixed for 24 hours, dehydrated in a graded series of alcohol solutions (70% to 100%), cleared with xylene, and embedded in paraffin. Cross-sections of the segments were cut at a thickness of approximately 5 μm with a microtome.c Sections were stained with H&E. Villus height, crypt depth, and villus width were evaluated in each intestinal segment. Villus height was defined as the distance from the villus tip to the crypt mouth, crypt depth was defined as the distance from the crypt mouth to the base, and villus width was defined as the widest distance of the villus. In each section, 10 villi and their associated crypts were measured via a light microscope with a computerassisted morphometric system.d The villus height-to-crypt depth ratio and villus surface area within each segment were calculated.
Measurement of enzyme-specific activity—Mucosal samples were weighed and suspended in ice-cold PBS-EDTA solution (0.05M Na3PO4, 2.0M NaCl, and 0.002M EDTA [pH, 7.4]) by use of a ratio of 1:10 (wt/vol). Samples were homogenized for 45 seconds and centrifuged at 2,000 X g for 10 minutes. The protein supernatant was collected from each sample, and protein concentration was measured via the Lowry method17 by use of a detergent-compatible protein assay,e with bovine serum albumin as the standards.
Activity of lactase, sucrase, and maltase was determined as described elsewhere,18 with lactose, maltose, and sucrose as the respective substrates. In addition, liberation of glucose was measured by use of a glucose kit.f Briefly, 100 μL of supernatant was added to a test tube and incubated with the respective substrate (ie, lactose, sucrose, or maltose; 100 μL; 0.056 mol/L) for 60 minutes at 37°C. Reactions were stopped by submerging the tubes in boiling water for 2 minutes. The tubes were allowed to cool, and glucose concentrations were determined.
Immunohistochemical staining for detection of PCNA, iNOS, and PPARγ—Slides with 5-μm-thick tissue sections were deparaffinized, rehydrated, and hydrated with buffer (PBS solution [pH, 7.4]). The slides were treated with 3% H2O2 in methanol for 10 minutes to quench endogenous peroxidase activity and heated in a microwave in 10mM citrate buffer (pH, 6.0) to retrieve antigen. Slides were washed, blocked by incubation with 5% bovine serum albumin in PBS solution for 30 minutes at 25°C, and then exposed to the primary antibodies (mouse monoclonal antibody against PCNAg [dilution, 1:100], rabbit polyclonal antibody against iNOSh [dilution, 1:50], and rabbit polyclonal antibody against PPARγi [dilution, 1:20]) by incubation overnight at 4°C. The antibodies against PCNA, iNOS, and PPARγ have been validated for use in pigs by Gookin et al,19 Wan et al,20 and Liu et al,6 respectively After incubation with the primary antibodies, slides were incubated for 30 minutes at 25°C in goat anti-mouse IgGj (dilution, 1:100) or goat anti-rabbit IgGk (dilution, 1:100), which was followed by incubation with avidin-biotinylated peroxidase complex for 20 minutes. The reaction was developed by staining with 3,3′-diaminobenzidine substratek; the reaction was stopped by immersion in distilled water, and slides were counterstained with hematoxylin for 25 seconds. Negative control sections were not treated with primary antibody.
For each slide, the number of cells that had positive results for staining with PCNA, iNOS, or PPARγ was counted in at least 10 fields. In each intestinal segment, PCNA-positive cells were counted in crypt cells; iNOS-positive cells were counted in blood vessels, mesenchyme, and villi; and PPARγ-positive cells were counted in epithelium and mesenchyme. The proliferation index, percentage of iNOS-positive cells, and percentage of PPARγ-positive cells were calculated as the ratio of the number of cells that had positive staining results for PCNA, iNOS, or PPARγ, respectively, to the total number of cells.
Photomicrographs of representative tissue sections obtained from an intestinal blood vessel (A), the intestinal mesenchyme (B), and an intestinal villus (C) in the duodenum of a pig after treatment with LPS. Cells that stained positively for iNOS are indicated (arrows). Tissue sections were incubated with rabbit polyclonal antibody against iNOS (primary antibody), followed by goat anti-rabbit IgG (secondary antibody), and then by avidin-biotinylated peroxidase complex. Bar = 86 μm.
Citation: American Journal of Veterinary Research 71, 11; 10.2460/ajvr.71.11.1331
Statistical analysis—All data were analyzed via an ANOVA appropriate for a completely randomized block design.l Results for pigs injected with LPS were compared by use of preplanned contrasts with results for control pigs to determine the effect of LPS administration or with results of pigs treated with rosiglitazone plus LPS to determine the effect of rosiglitazone in pigs challenge exposed to LPS. Results were reported as least square means ± pooled SEM. Significant differences were calculated by use of the least square means; values were considered significant at P < 0.05.
Results
Effects of LPS injection—Twenty minutes after LPS injection, all 12 pigs administered LPS had clinical signs of LPS-induced endotoxemia. Those signs included fever, vomiting, shivering, anorexia, and hypersomnia.
Intestinal morphology—In all 3 small intestinal segments, there was no difference in villus height among treatments (Table 1). The LPS-treated pigs had a significant increase in crypt depth in the duodenum (P = 0.001) and jejunum (P = 0.035) but not in the ileum (P = 0.089) and a significant decrease in the villus height-to-crypt depth ratio in the duodenum (P < 0.001), jejunum (P = 0.013), and ileum (P = 0.031), compared with results for the control pigs. Compared with results for LPS-treated pigs, pigs treated with rosiglitazone plus LPS had a significant decrease in crypt depth in the duodenum (P < 0.001) but not in the jejunum (P = 0.062) or ileum (P = 0.068) and a higher villus height-to-crypt depth ratio in the duodenum (P < 0.001), jejunum (p = 0.002), and ileum (P = 0.006). Treatment with rosiglitazone plus LPS had no effect on villus surface area in all 3 intestinal segments.
Effect of LPS and rosiglitazone plus LPS on intestinal morphology of weaned pigs (n = 6 pigs/treatment group).
| P value† | ||||||
|---|---|---|---|---|---|---|
| Variable | Control* | LPS* | Rosiglitazone plus LPS* | SEM | LPS vs control | LPS vs rosiglitazone plus LPS |
| Villus height (μm) | ||||||
| Duodenum | 280 | 275 | 279 | 23 | 0.827 | 0.848 |
| Jejunum | 289 | 289 | 307 | 22 | 0.988 | 0.443 |
| Ileum | 293 | 282 | 319 | 30 | 0.733 | 0.237 |
| Crypt depth (μm) | ||||||
| Duodenum | 94 | 165 | 88 | 16 | 0.001 | <0.001 |
| Jejunum | 123 | 162 | 128 | 17 | 0.035 | 0.062 |
| Ileum | 115 | 144 | 113 | 16 | 0.089 | 0.068 |
| Villus height-to-crypt depth ratio | ||||||
| Duodenum | 3.39 | 1.85 | 3.42 | 0.22 | <0.001 | <0.001 |
| Jejunum | 2.62 | 2.12 | 2.77 | 0.18 | 0.013 | 0.002 |
| Ileum | 2.98 | 2.26 | 3.24 | 0.30 | 0.031 | 0.006 |
| Villus width (μm) | ||||||
| Duodenum | 189 | 164 | 182 | 12 | 0.053 | 0.149 |
| Jejunum | 147 | 140 | 142 | 9 | 0.436 | 0.815 |
| Ileum | 152 | 166 | 159 | 15 | 0.370 | 0.661 |
| Villus surface area (μm2) | ||||||
| Duodenum | 166,695 | 143,170 | 161,441 | 20,770 | 0.275 | 0.393 |
| Jejunum | 133,139 | 127,358 | 137,440 | 13,489 | 0.674 | 0.466 |
| Ileum | 140,340 | 151,942 | 159,570 | 23,767 | 0.632 | 0.753 |
Control pigs were injected IP with DMSO and then injected IP 30 minutes later with sterile saline (0.9% NaCl) solution, LPS-treated pigs were injected IP with DMSO and then injected 30 minutes later with Escherichia coli LPS (100 μg/kg, IP), and rosiglitazone plus LPS-treated pigs were injected with rosiglitazone (3 mg/kg, IP) and then injected 30 minutes later with E coli LPS (100 μg/kg, IP).
*Values reported are least square means.
†Values were considered significant at P < 0.05.
Disaccharidase activity—Administration of LPS significantly (P = 0.006) decreased lactase activity in the duodenum, and it decreased (but not significantly) sucrase activity in the duodenum (P = 0.054) and lactase activity in the ileum (P = 0.075; Table 2). Rosiglitazone had no effect on enzyme activity in all 3 intestinal segments.
Effect of LPS and rosiglitazone plus LPS on disaccharidase activity in intestinal mucosa of weaned pigs (n = 6 pigs/treatment group).
| P value† | ||||||
|---|---|---|---|---|---|---|
| Variable | Control* | LPS* | Rosiglitazone plus LPS* | SEM | LPS vs control | LPS vs rosiglitazone plus LPS |
| Sucrase | ||||||
| Duodenum | 0.83 | 0.69 | 0.76 | 0.07 | 0.054 | 0.297 |
| Jejunum | 3.59 | 3.74 | 3.32 | 0.42 | 0.726 | 0.332 |
| Ileum | 4.14 | 4.99 | 5.34 | 0.56 | 0.153 | 0.535 |
| Lactase | ||||||
| Duodenum | 1.69 | 1.17 | 1.09 | 0.16 | 0.006 | 0.640 |
| Jejunum | 3.28 | 3.72 | 3.16 | 0.51 | 0.396 | 0.290 |
| Ileum | 0.86 | 0.60 | 0.64 | 0.14 | 0.075 | 0.782 |
| Maltase | ||||||
| Duodenum | 5.69 | 5.93 | 5.43 | 0.29 | 0.418 | 0.104 |
| Jejunum | 7.09 | 7.02 | 6.54 | 0.28 | 0.815 | 0.111 |
| Ileum | 8.18 | 7.55 | 8.05 | 0.39 | 0.126 | 0.215 |
*Values reported are least square mean U/mg of protein.
†Values were considered significant at P < 0.05.
See Table 1 for remainder of key.
Photomicrographs of representative tissue sections obtained from the intestinal mesenchyme (A) and the intestinal epithelium (B) in the duodenum of a pig after treatment with LPS. Cells that stained positively for PPARγ are indicated (arrows). Tissue sections were incubated with rabbit polyclonal antibody against PPARγ (primary antibody), followed by goat anti-rabbit IgG (secondary antibody), and then by avidin-biotinylated peroxidase complex. Bar = 86 μm.
Citation: American Journal of Veterinary Research 71, 11; 10.2460/ajvr.71.11.1331
Crypt cell proliferation—A significant decrease in the crypt cell proliferation index was observed in the jejunum (P < 0.001) and ileum (P < 0.001) but not in the duodenum (P = 0.072) of LPS-treated pigs, compared with results for control pigs (Table 3). Rosiglitazone administered 30 minutes before LPS administration significantly (P < 0.001) prevented the LPS-induced decrease of crypt cell proliferation in the jejunum and ileum (P < 0.001), compared with results for administration of DMSO followed by administration of LPS.
Effect of LPS and rosiglitazone plus LPS on crypt cell proliferation in intestinal mucosa of weaned pigs (n = 6 pigs/treatment group).
| P value† | ||||||
|---|---|---|---|---|---|---|
| Variable | Control* | LPS* | Rosiglitazone plus LPS* | SEM | LPS vs control | LPS vs rosiglitazone plus LPS |
| Duodenum | 46.27 | 41.59 | 42.89 | 2.42 | 0.072 | 0.599 |
| Jejunum | 45.37 | 39.53 | 46.79 | 1.29 | <0.001 | <0.001 |
| Ileum | 43.70 | 33.09 | 41.83 | 1.90 | <0.001 | <0.001 |
*Values reported are least square mean percentages.
†Values were considered significant at P < 0.05.
See Table 1 for remainder of key.
Percentage of iNOS-positive cells—In the 3 intestinal segments, cells staining positively for iNOS were observed mainly in vascular endothelial cells of blood vessels and, to a lesser extent, in fibroblasts of the mesenchyme and villi (Figure 1). Cells staining positively for iNOS were not observed in the crypts. The LPStreated pigs had a significant (P = 0.001) increase in iNOS staining in cells of the jejunal villi but not in cells of the duodenal mesenchyme (P = 0.065) or ileal blood vessels (P = 0.085), compared with results for the control pigs (Table 4). Compared with LPS-treated pigs, pigs treated with rosiglitazone plus LPS had a significant (P = 0.01) decrease in iNOS staining in cells of the duodenal blood vessels, villi, and ileal blood vessels but not in cells of the jejunal blood vessels (P = 0.091).
Effect of LPS and rosiglitazone plus LPS on the number of cells that stained positively for iNOS in intestinal mucosa of weaned pigs (n = 6 pigs/treatment group).
| P value† | ||||||
|---|---|---|---|---|---|---|
| Variable | Control* | LPS* | Rosiglitazone plus LPS* | SEM | LPS vs control | LPS vs rosiglitazone plus LPS |
| Duodenum | ||||||
| Blood vessels | 28.08 | 33.36 | 20.46 | 4.28 | 0.236 | 0.009 |
| Mesenchyme | 1.93 | 2.60 | 2.05 | .034 | 0.065 | 0.125 |
| Villi | 2.20 | 2.85 | 2.33 | 0.46 | 0.176 | 0.274 |
| Jejunum | ||||||
| Blood vessels | 37.74 | 39.47 | 30.38 | 5.03 | 0.735 | 0.091 |
| Mesenchyme | 2.32 | 2.43 | 2.21 | 0.55 | 0.854 | 0.698 |
| Villi | 2.20 | 5.34 | 2.52 | 0.77 | 0.001 | 0.002 |
| Ileum | ||||||
| Blood vessels | 33.62 | 41.49 | 28.96 | 4.26 | 0.085 | 0.010 |
| Mesenchyme | 2.25 | 2.92 | 2.49 | 0.52 | 0.218 | 0.425 |
| Villi | 4.10 | 5.98 | 5.62 | 1.08 | 0.104 | 0.746 |
*Values reported are least square mean percentages.
†Values were considered significant at P < 0.05.
See Table 1 for remainder of key.
Percentage of PPARγ-positive cells—In all 3 intestinal segments, cells staining positively for PPARγ were observed mainly in vascular endothelial cells and lymphocytes of the mesenchyme and, to a lesser extent, in epithelial cells of the epithelium (Figure 2). Compared with control pigs, LPS-treated pigs had a significant decrease in the percentage of PPARγ-positive cells in the duodenal (P = 0.018) and ileal (P = 0.041) epithelium and the jejunal (P = 0.001) and ileal (P = 0.01) mesenchyme (Table 5). Compared with LPS-treated pigs, pigs treated with rosiglitazone followed by LPS had a significant increase in PPARγ staining in the duodenal (P < 0.001), jejunal (P = 0.003), and ileal (P < 0.001) mesenchyme but not in the ileal epithelium (P = 0.057).
Effect of LPS and rosiglitazone plus LPS on the number of cells that stained positively for PPAR? in intestinal mucosa of weaned pigs (n = 6 pigs/treatment group).
| P value† | ||||||
|---|---|---|---|---|---|---|
| Variable | Control* | LPS* | Rosiglitazone plus LPS* | SEM | LPS vs control | LPS vs rosiglitazone plus LPS |
| Duodenum | ||||||
| Epithelium | 6.28 | 4.75 | 5.08 | 0.58 | 0.018 | 0.575 |
| Mesenchyme | 19.91 | 20.56 | 26.54 | 1.28 | 0.622> | <0.001 |
| Jejunum | ||||||
| Epithelium | 5.16 | 5.27 | 4.77 | 0.74 | 0.890 | 0.516 |
| Mesenchyme | 18.30 | 13.80 | 17.96 | 1.15 | 0.001 | 0.003 |
| Ileum | ||||||
| Epithelium | 9.38 | 7.68 | 9.25 | 0.76 | 0.041 | 0.057 |
| Mesenchyme | 19.49 | 15.42 | 24.78 | 1.36 | 0.009 | <0.001 |
*Values reported are least square mean percentages.
†Values were considered significant at P < 0.05.
See Table 1 for remainder of key.
Discussion
In the study reported here, injection of LPS was used to induce intestinal injury in pigs, which enabled us to evaluate whether rosiglitazone, a PPARγ ligand, could attenuate intestinal injury in weaned pigs. Lipopolysaccharide is a component of the cell wall of gram-negative bacteria. It is generally accepted that LPS induces signs of illness, including respiratory failure, anorexia, diarrhea, somnolence, and hypersomnia.6,21 Lipopolysaccharide can also cause various morphologic changes in the intestines, such as villus atrophy, epithelial vacuolation, submucosal edema, frank hemorrhage, and necrosis.3,22,16
Villus height and crypt depth are often considered as indices that reflect intestinal morphology.3 In the present study, injection of LPS caused intestinal injury in weaned pigs, which was indicated by an increase in crypt depth and a decrease in the villus height-to-crypt depth ratio. Similarly, investigators in another study16 reported that severe damage to the intestinal mucosa was evident in pigs at 3 hours after LPS challenge exposure. Compared with the LPS-treated pigs, pigs treated with rosiglitazone plus LPS had an attenuation of the increase in crypt depth and the decrease in the villus height-to-crypt depth ratio caused by administration of LPS. Analysis of these data indicated that rosiglitazone protected the intestinal mucosa from damage. In agreement with the results of our study, treatment with rosiglitazone reduced the score for macroscopic damage to the colon mucosa and the morphologic alterations associated with acute TNBS administration in 1 study9 and reduced the severity and extension of chronic colonic damage induced by TNBS in another study.23 In addition, rosiglitazone has been found to decrease ischemia-reperfusion injury of the intestines.24
Activities of mucosal disaccharidases, including sucrase, lactase, and maltase, have been used as indicators of mucosal maturation and digestive function of the intestines in pigs.25 In the study reported here, LPS decreased sucrase activity in the duodenum and lactase activity in the duodenum and ileum. However, rosiglitazone did not alleviate the decrease in disaccharidase activity induced by LPS administration, which indicated that rosiglitazone was not effective for improving digestive function.
Integrity of the intestinal mucosa depends on the continued proliferation, migration, and differentiation of enterocytes in the intestinal crypts.3,26 Impairment of enterocyte proliferation in the intestinal crypts may be a key factor that contributes to mucosal injury and failure of the intestinal barrier. In the present study, LPS administration decreased the crypt cell proliferation index in weaned pigs, which is in agreement with results in rats.27,28 Rosiglitazone alleviated the decrease in the crypt cell proliferation index induced by LPS administration. The PPARγ agonists, such as rosiglitazone, can exert an antiproliferation effect in cancer cells, such as MCF-7 breast cancer cells,29 colon cancer cells,30 and pancreatic cancer cells.31 Few studies have been conducted to examine the effects of PPARγ agonists on proliferation of normal intestinal epithelial cells. Analysis of data for the study reported here indicated that rosiglitazone treatment might alleviate intestinal mucosa injury via prevention of the LPS-induced decrease in crypt cell proliferation.
The PPARγ agonists can attenuate the severity of inflammation in rodents and humans.8,32,33 The local increase in iNOS expression correlates with an increase in the severity of intestinal injury.34 In the present study, iNOS was expressed mainly in intestinal vascular endothelial cells. Vascular endothelial cells play a key role in inflammation by undergoing activation and recruiting circulating immune cells into tissues and foci of inflammation, which is an early and rate-limiting step in the inflammatory process.35 Several conditions, such as intestinal inflammation and sepsis, are related to overexpression of iNOS in macrophages and endothelial cells.36 In the study reported here, treatment with rosiglitazone caused an attenuation of inflammation in the intestines, as reflected by a reduction of iNOS expression. The inhibitory effect of rosiglitazone on intestinal inflammation in this study is similar to that reported in other studies.15,37 In fact, rosiglitazone can attenuate the increase in iNOS activity in the brain of stressed rats.37 Additionally, investigators in 1 study15 reported that rosiglitazone attenuated the expression of iNOS and cyclooxygenase-2 in carrageenan-induced paw edema and carrageenan-induced pleurisy of rats.
Induction of iNOS may be responsible for tissue injury via the formation of nitric oxide–dependent nitrating species, such as peroxynitrite. Overproduction of nitric oxide or its toxic metabolite, peroxynitrite, after endotoxemia promotes and sustains mucosal injury and failure of the intestinal barrier, in part, by inhibiting enterocyte proliferation.38 In the present study, an increase in expression of iNOS was accompanied by a decrease in crypt cell proliferation. Thus, rosiglitazone may prevent LPS-induced decrease of crypt cell proliferation through a decrease in iNOS expression to alleviate intestinal damage.
The PPARγ agonists may exert their anti-inflammatory effects by negatively regulating the expression of proinflammatory genes (ie, iNOS) via PPARγ-dependent or -independent mechanisms. To investigate whether the observed beneficial effects of rosiglitazone on the intestines were mediated through a PPARγ-dependent pathway, we also determined the percentage of PPARγ-positive cells in the intestines. Analysis of the data indicated that LPS administration decreased the percentage of PPARγ-positive cells in the duodenum, jejunum, and ileum at 3 hours after LPS injection. However, in another study3 conducted by our laboratory group, we found that LPS injection increased PPARγ mRNA content in the jejunum at 6 hours after LPS administration. The reasons for this discrepancy might be that the effects of LPS administration on PPARγ expression in the intestines may be a time-dependent event. In that study3 and the study reported here, different time points (3 hours in the present and 6 hours in our previous study) were used for the collection of intestinal samples. Similar to results of the present study, PPARγ expression is downregulated in a number of inflammatory bowel diseases, such as experimentally induced colitis,12 ulcerative colitis,34 and Crohn's disease.39 In addition, we detected that the percentage of PPARγ-positive cells in the intestines increased after rosiglitazone treatment. Similar to results of the present study, rosiglitazone has been found to activate PPARγ in other studies.40,41 Rosiglitazone induced PPARγ transcriptional activity in human adrenocortical carcinoma H295R cells.40 In another study,41 investigators reported that the protective effects of rosiglitazone in experimentally induced lung injury are dependent on activation of PPARγ The fact that rosiglitazone treatment resulted in an increase in the percentage of PPARγ-positive cells suggested that PPARγ signaling might be activated. Therefore, we speculated that the modulatory effect of rosiglitazone on the synthesis of proinflammatory mediators (eg, iNOS) might be through a PPARγ-dependent mechanism. Regretfully, PPARγ transcriptional activity was not measured in the present study. In agreement with our speculation, investigators in another study15 reported that rosiglitazone attenuated the expression of iNOS and cyclooxygenase-2 in carrageenan-induced paw edema and carrageenan-induced pleurisy of rats via a PPARγ-dependent pathway. To evaluate these mechanisms, additional studies are needed that would include a PPARγ antagonist (such as bisphenol A diglycidyl ether15 or GW966242) in the experimental designs.
The proposed mechanism for the effect of PPARγ agonists (eg, rosiglitazone) on inflammation may be in suppression of the activation of inflammatory genes (eg, iNOS) by negatively interfering with nuclear factor-kB, signal transducers and activators of transcription-1, and activator protein-1 signaling pathways in a DNA binding-independent manner.4 Downstream of these transcription factors, pathways include the major proinflammatory cytokines, cell-adhesion molecules, and iNOS.4
In the study reported here, treatment with rosiglitazone exerted beneficial effects in alleviating mucosal injury in the intestines of LPS-treated pigs. The protective effects of rosiglitazone on the intestines may be associated with a decrease in the expression of intestinal proinflammatory mediators (eg, iNOS).
Abbreviations
| DMSO | Dimethyl sulfoxide |
| iNOS | Inducible nitric oxide synthase |
| LPS | Lipopolysaccharide |
| PCNA | Proliferating cell nuclear antigen |
| PPAR | Peroxisome proliferator-activated receptor |
| TNBS | 2,4,6-trinitribenzene sulfonic acid |
Sigma Chemical Co, St Louis, Mo.
Alexis, Milan, Italy.
American Optical Co, Scientific Instrument Division, Buffalo, NY.
BioScan Optimetric, BioScan Inc, Edmonds, Wash.
Bio-Rad Laboratories, Hercules, Calif.
Sigma Diagnostics, St Louis, Mo.
PC10, Dako Ltd, High Wycombe, Buckinghamshire, England.
Transduction Laboratories, Lexington, Ky.
Abcam, Cambridge, England.
Boster Biotech, Wuhan, China.
Dako Laboratories, Carpinteria, Calif.
PROC GLM, SAS, version 8.2, SAS Institute Inc, Cary, NC.
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