The Effect of Short-chain Fatty Acid Supplementation on Acute Colonic Inflammation in Rats Fed With Western Diet: Role of the Peroxisome Proliferator-activated Receptor-gamma

doi:10.21203/rs.3.rs-1067920/v1

Increased incidence of inflammatory bowel disease (IBD) is associated with the consumption of westernized diet (WD). In male Wistar albino rats induced with colitis by oral intake of dextran-sulfate sodium (DSS) for 10 days, we investigated how colitis-induced oxidative damage is affected by WD consumption started 3 weeks before DSS intake. Secondly, the putative protective effects of short-chain fatty acids (SCFAs) were evaluated in both WD-fed and normal diet (ND)-fed rats. WD consumption prior to colitis induction alleviated oxidative damage and suppressed inflammatory response of the colon via the modulation of peroxisome proliferator-activated receptor (PPAR)-γ activity, suggesting a preconditioning impact of the WD. Moreover, addition of SCFA to WD abolished the elevations in lipid profile, while SCFA added to either ND or WD during the 10-day development of colonic inflammation depressed the pro-inflammatory TNF-α and IL-6 expressions and upregulated IL-10, but the reduction in oxidative damage due to WD was not further changed. The results demonstrated that acute colonic inflammation is alleviated when it is preceded by the consumption of WD, and SCFAs improved WD-induced disruption in lipid profile, implicating that SCFAs would be advantageous in IBD patients comorbid with metabolic syndrome.

Physiology

westernized diet

bowel inflammation

short-chain fatty acids

oxidative damage

peroxisome proliferator-activated receptor gamma

Inflammatory bowel disease (IBD) is a chronic disease characterized by the deterioration of the mucosal balance, which involves the overgrowth of bacteria in the gut along with changes in the synthesis of cytokines and inflammatory mediators [1]. Westernized diet (WD), which typically contains less fruits and vegetables but more fat, sugar and processed meat [2], has been also proposed as an important reason for the increased incidence of IBD especially in the developing countries [2-4]. Degradation of the intestinal epithelial barrier followed by the activation of immune system is considered as the primary mechanism in the development of IBD [5]. Increased consumption of fat and sugar and decreased consumption of vegetables and fiber alter the composition of the gut microbiota and make the host mucosa open to pathogens [4, 6], while enhanced production of the pro-inflammatory cytokines promotes inflammatory processes in the intestines [7, 8]. Activated immune responses in the gut involve the overproduction of pro-inflammatory cytokines, such as tumour necrosis factor alpha (TNF-α), interleukin (IL)-1 beta (IL-1β), IL-6, IL-8, and interferon gamma (IFN-γ), which enhance the inflammatory cascades by triggering the formation of other pro-inflammatory elements, and collecting neutrophils to the site of mucosal injury [9]. The recruitment of neutrophils to the inflamed tissue exaggerates the mucosal damage and yields to apoptosis by releasing reactive oxygen species (ROS) [5, 10], which further increase the production of several inflammatory mediators by activating nuclear factor-κB (NF-κB) [11].

Short-chain fatty acids (SCFAs), the saturated fatty acids with less than six carbon atoms, are the end products of fermentation produced by the intestinal microbiota via the utilization of some non-digestible carbohydrates [12]. In the human large intestine, members of the Bacteroidetes phylum mainly produce acetate and propionate, while Firmicutes mostly produces butyrate [13, 14], and the amount and relative proportion of each SCFA depend on the substrate, microbiota composition and gut transit time [12, 15, 16]. SCFAs were shown to have various effects on colonic health, particularly on the maintenance of epithelial lining by modulating the turnover and differentiation of colonic epithelial cells. When luminal substrates were reduced by parenteral nutrition, elemental diet or by the diversion of the faecal stream, mucosal hypoplasia and atrophy were observed in experimental animals and humans [17-19]. Moreover, owing to its modulatory effects on the composition of the mucous layer and oxidative stress, low concentrations of butyrate were associated with the occurrence of several colonic diseases [17]. On the other hand, supplying with SCFAs was suggested to exert an anti-tumour effect on colonic carcinoma cell lines [20, 21], while SCFAs have an anti-inflammatory activity on colonic epithelial cells by inhibiting NF-κB [22, 23]. Accordingly, in vitro studies have indicated that SCFAs regulate the expression of several inflammatory cytokines (e.g. IL‐8, IL‐17, IL‐1β, IL‐6, IL‐12 and TNF‐α) in the colonic epithelial cells [24, 25].

Apart from the experiments related with colonic diseases, SCFAs were also shown to modulate the inflammatory processes induced by metabolic diseases [26]. Although studies have shown separately that SCFAs could improve various inflammatory disorders as well as the inflammation of the bowel [3, 22], the effect of SCFAs on inflammatory processes of colitis when it is accompanied by WD consumption, which widely occurs in developing countries, was not studied to date. In the present study, we firstly aimed to investigate how western diet counteracts with bowel inflammation in rats induced with colitis following a 3-week consumption of WD. Secondly, the putative protective effects of SCFAs were evaluated in both WD-fed and normal diet-fed rats with colitis.

2.1. Ethical approval

All the experimental procedures and the design of the study were approved by the Marmara University Animal Care and Use Committee (Date: 5 November 2018; No: 97.2018.mar). The experiments were performed in compliance with the Turkish law on the use of animals in experiments, regarding the ARRIVE guidelines and the guidelines of the New York Academy of Sciences.

2.2. Animals

A total of forty Wistar albino male rats (200-300 g), obtained from the Marmara University Animal Centre, were kept in 12-hour light-dark cycles and at a room temperature of 22-24°C. Four rats were housed in a cage with free access to water and rat chows (normal diet; ND or WD), which were delivered daily in portions of 100 g. During the 38-day follow-up period of the experiment, body weights of the animals were measured daily.

2.3. Experimental design

After a 7-day adaptation, rats were randomly divided into one of the feeding regimens, and either ND (58% carbohydrate, 18% fat and 24% protein) or WD (36% carbohydrate, 45% fat and 19% protein) was supplied for the following 31 days [27, 28]. Except for the control group, which received ND and tap water, all the other groups were given dextran sodium sulphate (DSS) in their drinking water to induce colonic inflammation during the last 10 days of the follow-up period (Fig. 1). Oral administration of DSS is a practical and reproducible method to induce disruption of the colonic epithelial barrier, and it is commonly used as an animal model of colonic inflammation that mimics IBD [29-31]. Accordingly, DSS solution (0.5%) was freshly prepared every day [31] and was given to four groups of animals for 10 days in their drinking water. A mixture of SCFAs (150 mM/L; 60% acetate, 20% propionate and 20% butyrate) was freshly prepared and added to the DSS-containing drinking water of a ND-fed and a WD-fed group [32, 33], while DSS was given alone in tap water in the remaining ND-fed and WD-fed groups. All applications were carried out between 8:30 to 12:00 AM. Body weights of the rats were recorded every other day throughout the follow-up period. At the end of the 31 days, colonic blood flow measurements were made under anesthesia and then serum, faeces, liver and colon samples were obtained. The colon was opened longitudinally and cleared with saline. Macroscopic scoring of the colonic damage was performed by one researcher who was unaware of the treatment groups, using the following criteria: intestinal consistency (0-2), intestinal colour (0-2), and intestinal dilatation (0-2) with a maximum score of 6 [34].

2.4. Colonic blood flow measurement

Under ketamine (100mg/kg; intraperitoneally, ip) and xylazine (10mg/kg, ip) anesthesia, transverse colon was excised following a midline laparotomy. Blood flow was measured using Doppler ultrasound device (PeriFlux system 5000, PERIMED, Sweden, 2010) and PeriSoft 2.5.5 software). The probe (Probe 307) was placed on the serosal surface of the colon and a 1-min recording was made. Three ten-second episodes with at least 7-sec intervals were selected from each recording and the average perfusion unit (PU) was calculated. Immediately after the blood flow measurements, the rats were euthanized by cardiac puncture.

2.5. Measurement of the parameters in the serum

Serum levels of triglycerides (TG), total cholesterol (TC), low-density lipoprotein (LDL), aspartate aminotransferase (AST), alanine aminotransferase (ALT), albumin and glucose were measured by spectrophotometric methods using a AU5800 automatic biochemical analyser (Beckman Instruments, Fullerton, CA, USA).

2.6. Myeloperoxidase activity (MPO) measurements in the colon and liver samples

Myeloperoxidase (MPO), an enzyme found principally in the azurophilic granules of polymorphonuclear leukocytes (PMNL), is frequently used to estimate PMNL accumulation in the inflamed tissues, and shows a positive correlation with the histochemically-determined PMNL abundance in the tissues. For this purpose, after washing tissues with phosphate buffered saline (PBS), tissue samples of colon and liver were homogenized by a detergent buffer (in 50 mM potassium phosphate, 0.5% HETAB, pH 6.0, 10 mM EDTA), then centrifuged at 10 min, at 4°C 12000 rpm. After the supernatant was removed, the pellet was homogenized again in a 50 mM potassium-phosphate buffer. MPO activity was determined by spectrophotometric measurement of the 2HCl (20 mg/ml) - dependent oxidation of o-dianizidine to H₂O₂^-(20 mM) at 460 nm and MPO activity was expressed as U/g [35, 36].

2.7. Glutathione (GSH) determination in the colon and liver samples

Glutathione (GSH) is an antioxidant synthesized in the cells to defend against the oxidant damage. Tissue samples were homogenized with a 10% solution of trichloric acid equivalent to 10 times of the weight of the tissue, and were centrifuged at 3000 rpm, 4°C for 15 minutes. In glutathione measurements, the modified Ellman method was used, and the amount of GSH was determined spectrophotometrically. Briefly, after centrifugation 250 µl of supernatant was added to 1 mL of 0.3 mol/L Na₂HPO₄·H₂O. Then, 125 µl solution of 5,5'-dithiobis(2-nitrobenzoic acid) (0.4 mg/mL in 1% sodium citrate solution) was added to this mixture.2H2O solution. A 0.2-mL solution of dithiobisnitrobenzoate (0.4 mg/mL 1% sodium citrate) was added and the absorbance at 412 nm was measured immediately after mixing. The amount of GSH was read in 412 nm absorbance in a spectrophotometer, and GSH was expressed in µmol/g/tissue [35, 37, 38].

2.8. Chemiluminescence measurements in the colon and liver samples

Chemiluminescence is commonly utilized as a direct and non-invasive method for the measurement of ROS by luminol and lucigenin probes, which produce a highly stimulated product when added to in vitro organic systems, and the resultant light energy (chemiluminescence) can be measured by a luminometer . Lucigenin, selective to O₂^-(0.2 mM; bis-N-methylacridinium nitrate) or luminol (5-Amino-2,3-dihydro-1,4-phthalazinedione), which detects radicals such as OH^-, H₂^-O₂^- and HOCl, was added in a 0.2 mM concentration, and measurements were made with luminometer at room temperature (Junior LB 9509 luminometer; EG&G Berthold). Counts were obtained at 1-min intervals, and the results are given as the area under the curve (AUC) for a counting period of 5 minutes. The data are expressed as AUC after the numbers are verified by the tissue weights. The results are expressed as relative light unit /mg tissue (rlu/mg) [39].

2.9. Examination of bacterial colony-forming units (CFU) in the fecal and hepatic samples

The liver and fecal samples were collected aseptically following cardiac puncture. Liver samples were excised, cut into small pieces and transferred into sterile micro-tubes and dry weighed. Phosphate-buffered saline (PBS, pH:7.4) was added to the tubes to obtain a concentration of 1 g/mL and mixed vigorously using vortex. A 0,1 mL homogenate was serially diluted 10^-1 to 10^-4 mL in PBS. Hundred mL samples were taken and inoculated on blood agar media (bioMérieux, Marcy l'Etoile, France). Plates were incubated in a lab incubator (Memmert GmbH, Schwabach, Germany) at 37°C for 24 hours. Resulting colonies were counted and expressed as CFU/mL. Experiments were performed in triplicates and the average was taken for each rat specimen.

2.10. Western blotting for the determination of cytokines in the colonic tissue

Western blot analyses were performed as previously described [40]. The bicinchoninic acid assay (BCA) was used to detect protein concentrations in homogenized samples. Afterwards, 25 µg of protein was resolved in 15% sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to polyvinylidene difluoride (PVDF) membrane (sc-3718, Santa Cruz Biotechnology), which was blocked with 3% bovine serum albumin (BSA) in Tris-buffered saline (TBS), washed twice in TBS containing 0.1% Tween-20 and incubated overnight with primary antibody (1:500 dilution monoclonal antibody anti-TNF-α sc-1351, anti-IL-6 sc-7920, anti-IL-10 sc-32815, anti-GAPDH sc-25778, Santa Cruz Biotechnology, Heidelberg, Germany). The membrane was incubated with horseradish peroxidase (HRP)-conjugated secondary antibody (1:2000 anti-mouse IgG1-HRP, sc-2060 and anti-rabbit IgG-HRP sc-2004, Santa Cruz Biotechnology) for two hours. Chemiluminescence reagents (sc-2048, Santa Cruz Biotechnology) using a Chemiluminescent Imaging System (Syngene, USA) were used to detect the blots. Data were analyzed using the ImageJ OD analysis software. Signals were normalized with respect to β-actin.

2.11. Western blotting for the determination of peroxisome proliferator-activated receptor (PPAR) gamma in the colonic tissue

Protein expressions of PPAR gamma 1-2 were determined by the western blotting method. The obtained results, used as a loading control, were compared to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) bands. According to the calculated optical densities, quantitative results were obtained. Tissues were lysed with Tris-HCl, NP-40, Sodium dodecyl sulfate and Pefabloc containing lysis solution and centrifuged. Following the denaturation of the obtained supernatant proteins, they were loaded onto 10% polyacrylamide gels and transferred to a nitrocellulose membrane. Using anti-PPAR gamma antibody (07-466, Merck Millipore, Darmstadt, Germany), protein expressions were determined according to the instructions of manufacturer.

2.12. Histopathological examination of the colonic tissue

Tissue samples taken from the colon were placed in 10% formaldehyde and routinely processed by embedding them in paraffin. Tissue sections (5 mm) were stained with hematoxylin and eosin (H&E) and examined under a photomicroscope (Olympus bx51, Tokyo, Japan). The microscopic scoring was done by an experienced histologist, who was unaware of the experimental groups. The histological score was determined by scoring epithelial damage/shedding, inflammatory cell infiltration, vascular congestion, and submucosal edema as 0=never, 1=mild, 2=medium, 3=severe (maximum score is 12) [35].

2.13. Justification of sample size and statistical analysis

The resource equation method was used to determine the sample size [41, 42]. The formula (20/k) + 1 (k: number of groups) [43, 44] was used and there are a total of 5 groups in the study. Since the small sample size may increase the possibility of false negatives and the study is an exploratory study, 1 was added to the sample size (5 + 1 = 6). However, as some data was previously predicted to be non-parametric, the sample size was increased by approximately 1/3 [45]. Finally, 8 animals per group were found to be sufficient for each group (8 x 5 = 40 animals in total).

All the statistical analyses were performed using GraphPad Prism 9.0 (GraphPad Software, San Diego, CA, USA). The data are expressed as mean ± standard error of the mean. Comparisons among different groups were analyzed by Kruskal-Wallis test followed by Post-hoc Dunn’s test (colon macroscopic scoring), one way or two-way ANOVA followed by post-hoc Tukey multiple comparison tests. Values of P < 0.05 were regarded as significant.

3.1. Effect of WD on DSS-induced colonic and hepatic inflammation

When compared to the colonic tissues of the control group, induction of colonic inflammation by oral administration of DSS for 10 days resulted in elevated blood flow (p<0.01), higher macroscopic score (p<0.001), increased neutrophil infiltration and enhanced production of ROS as detected by higher levels of luminol (p<0.01) and lucigenin (p<0.05) chemiluminescence (CL) (Fig. 2). In previously WD-fed rats, DSS-induced elevations in colonic blood flow, macroscopic score, MPO activity and lucigenin-enhanced CL levels were significantly depressed (p<0.05-0.001), suggesting a preventive impact of WD on a forthcoming inflammation of the bowel. Colonic and hepatic levels of GSH, an endogenous antioxidant, were not altered by neither DSS nor WD-feeding. However, neutrophil recruitment to the hepatic tissue was elevated in WD-fed rats with colonic inflammation (p<0.01), but MPO activity in the ND-fed rats was not different than that of the control group. Despite an observed tendency to increase, luminol- or lucigenin-detected ROS generation in the livers of DSS-administered rats was not significantly altered.

In parallel with these findings, protein expression levels of the pro-inflammatory cytokines IL-6 and TNF-α were significantly increased in the colonic tissues of ND-fed rats induced with DSS (p<0.001), while the expression of the anti-inflammatory IL-10 was depressed as compared to control rats (p<0.001; Fig. 3). Similar changes were observed in the WD-fed rats, but elevations in the IL-6 and TNF-α, as well as the decrease in IL-10 were significantly less in WD-fed rats with colonic inflammation (p<0.05-0.001).

3.2. Effect of WD on DSS-induced changes in the colonic protein expression of PPAR-γ

Induction of colonic inflammation in ND-fed rats did not significantly change cytoplasmic expression of PPAR- γ1 or PPAR- γ2 (Fig. 4). Despite any changes observed in the nuclear expression of PPAR- γ1, colonic inflammation resulted in an increased expression of nuclear PPAR- γ 2 in both the WD- and ND-fed groups (p<0.001), while the expression level was relatively lower in the WD-given group (p<0.001).

3.3. Effect of SCFA supplementation on DSS-induced inflammation of the colon and liver and on colonic PPAR- γ expression

In ND-fed rats induced with colonic inflammation, intake of SCFA significantly reduced colonic blood flow (p<0.001), but had no further effect on already depressed blood flow in the WD-fed rats (Fig. 5). On the other hand, supplementation with SCFA did not alter macroscopic scoring, MPO activity, GSH content or ROS generation in the colonic tissues of neither ND-fed nor WD-fed rats as compared to respective non-supplemented groups. In the liver tissues of ND-fed rats with colitis, SCFA increased GSH content (p<0.05) and inhibited the level of superoxide generation (p<0.01) as detected by lucigenin probe; but this protective effect of SCFA on hepatic oxidative damage was not present in the previously WD-fed rats. Supplementing the rats with SCFA during the colonic inflammatory process significantly reduced the protein expression levels of the pro-inflammatory cytokines, IL-6 and TNF-α, and elevated the anti-inflammatory IL-10 level in the colonic tissues of both ND- and WD-fed groups (p<0.001) (Fig. 3), demonstrating a diet-independent anti-inflammatory impact of SCFA on bowel inflammation. The upregulated colonic PPAR- γ expression in the ND- and WD-fed colitis groups was not further affected by supplementing with SCFA (Fig. 4).

3.4. Metabolic effects of colonic inflammation, WD feeding and SCFA supplementation

Despite a tendency to increase in the SCFA plus WD given group (11.5±1.6 %), the body weight changes during the follow-up period of the colitis groups fed with ND (7.3±3.2 %) or WD (5.1±4.6 %) or SCFA plus ND (8.1±1.6%) were not different than that of the control group (4.6±2.9%). The calculated daily calorie intake and liver wet weights of the animals (corrected to body weights) were also similar in all the experimental groups (data not shown). However, induction of colonic inflammation by DSS significantly increased serum glucose levels of both ND- and WD-given rats (p<0.05) (Table 1), but serum levels of TG and VDL levels were increased only in the WD-fed colitis group (p<0.001; Table 1). Addition of SCFA to WD did not change glucose levels, but abolished the elevations in TG and VLDL, showing an inhibitory effect of SCFA on WD-promoted disruption of the lipid metabolism. Moreover, feeding with WD or supplementing with SCFA in ND- or WD-given groups significantly elevated serum total albumin level (p<0.01-0.001). Despite the enhanced inflammatory response of the liver to colitis induction and to WD-feeding, which resulted in elevated MPO activity (Fig. 2), hepatic functions were not disturbed by DSS and/or WD (Table 1). However, addition of SCFA to diet elevated AST level in both ND- and WD-fed colitis groups, showing a negative impact of SCFA on hepatic function.

3.5. Effects of colonic inflammation, WD feeding and SCFA supplementation on microscopic damage and bacterial count in the colon and liver

Histological scoring of the colonic tissues revealed that colitis induction increased the microscopic damage and elevated the score (p<0.01), while the change in diet with or without the addition of SCFA did not have any further effect (p<0.05-0.01) (Fig. 6; Table 1). Bacterial count in the feces of DSS-administered rats was significantly reduced in both WD-fed (p<0.05) and ND-fed (p<0.01) rats (Table 1). On the other hand, DSS-induced significant reduction in fecal bacteria (p<0.05) was not altered by the addition of SCFA to ND or WD. Bacterial counts in the hepatic tissues of colitis-induced rats on either ND or WD were similar to that of the control group. However, when SCFA was added to WD, bacterial count in the liver tissue was increased significantly (p<0.01), showing an enhancement of colitis-induced bacterial translocation with the co-intake of SCFA and WD. On the other hand, the elevation in hepatic count of the bacteria has not reached to statistical significance when SCFA was added to ND.

Table 1. Serum levels of glucose, total albumin, triglycerides, VLDL, aspartate aminotransferase (AST), alanine aminotransferase (ALT); bacterial counts (colony forming units, CFU) in the colonic and hepatic tissues; and the histological damage scores of the colonic tissues of the experimental groups.

Parameters		Control		DSS
		Control		Normal diet		Western diet		Normal diet + SCFA		Western diet + SCFA
		Mean	SEM	Mean	SEM	Mean	SEM	Mean	SEM	Mean	SEM
Serum	Glucose (mg/dL)	214.6	19.7	300.8*	24.5	299.1^*	7.1	266.4	13.8	298.1	10.5
	ALT (U/L)	54.4	4.7	55.8	5.0	*38.8^,⁺**	2.1	54.5	3.0	51.75^#	3.8
	AST (U/L)	117.8	4.0	103.6	3.3	103.8	5.9	133.4^##	7.2	134.5^##	6.7
	Total albumin (g/L)	27.8	0.2	28.8	0.52	31.2^{,⁺⁺⁺}*	0.3	30.2	0.4	32.2	0.6
	Triglycerides	39.2	2.7	44.4	5.1	194.9^***	25.2	32.5	1.9	54.6	10.7
	VLDL	7.7	0.5	8.9	0.9	39.0^***	5.1	6.6	0.3	10.9	2.1
Bacterial count	Feces (CFU/mL)	17.0 x10¹¹	0.4x10¹¹	9.3x10¹⁰**	2.3x10¹⁰	21.6x10¹⁰*	8.7x10¹⁰	5.0 x10¹¹	1.5x10¹¹	7.7 x10¹¹	2.6x10¹¹
Bacterial count	Liver (CFU/mL)	15.7x10³	6.5x10³	8.1 x10³	2.1x10³	5.4x10³	2.1x10³	34.1 x10³	8.1 x10³	69.0x10^3#	26.2x10³
Histological scoring of colon		0.3	0.3	1.5**	0.3	1.3**	0.3	2.2	0.8	2.4	0.3

Data are expressed as mean ± SEM. *p<0.05, **p<0.01, ***p<0.001, compared to control group; +p<0.05, ++p<0.01, +++p<0.001, compared to normal diet- given DSS group; #p<0.05, ##p<0.01, ###p<0.001, compared to respective diet group that has not received SCFA. Ordinary one-way ANOVA followed by post-hoc Tukey test or ordinary two-way ANOVA followed by Post-hoc Tukey test was used.

Several studies have suggested that the dramatic increase in the incidence of IBD is due to westernized diet habits [7, 8], which include increased consumption of highly industrialized food [46, 47]. On the other hand, advantageous effects of SCFA on inflammation, including metabolic disorders and IBD were previously demonstrated [3, 22]. Based on these reports, the present study was aimed to explore specifically the impact of WD on the severity of the forthcoming colonic inflammation and the effect of short-term SCFA administration on the development of colonic inflammation. The current findings demonstrate that being on WD for three weeks prior to colitis induction abolished the increase in colonic blood flow, alleviated oxidative damage and suppressed the inflammatory response of the colon. Addition of SCFA to either ND or WD during the development of colonic inflammation depressed the pro-inflammatory cytokine expression and upregulated IL-10, but the reduction in oxidative damage due to WD was not further changed. Thus, DSS-induced oxidative damage of the colon was prevented with prior WD-feeding, while the presence of SCFA in the diet during the course of colitis downregulated the inflammatory response of the colon, but SCFA had no further effect on the anti-oxidative protection provided by WD. However, addition of SCFA to ND reduced hepatic ROS generation, elevated hepatic GSH content and reversed the WD-induced elevations in TG and VLDL, while colitis-induced increase in blood flow was abolished. Despite that colitis alone did not result in an apparent bacterial translocation, intake of SCFA in addition to WD caused a significant increase in bacterial colonization in the liver, suggesting the exaggerated exposure of the liver to intestinal microbes with the co-intake of SCFA and WD in the presence of bowel inflammation.

Clinical studies have revealed that IBD patients, independent of steroid use, have a relatively greater risk of diabetes [48] while they present with lower serum total cholesterol and TG levels [49]. Our findings showed that colitis induction, even in rats receiving ND, resulted in elevated levels of glucose, and as expected [50, 51] these changes were accompanied by increased TG and VLDL levels when the diet was previously shifted to WD, confirming that WD-feeding was able to induce metabolic alterations similar to those found in humans. On the other hand, administration of SCFA abolished the disrupted lipid profile in both colitis groups fed with either ND or WD. Previously, it was reported that dietary supplementation of SCFA improves the lipid profile of rodents fed with high-fat diet [52, 53], and in obese patients [54]. In support of these studies, we verified that SCFA improved lipid metabolism, which was altered by either diet or colonic inflammation. However, despite these metabolic disturbances, glucose levels, daily caloric intake and body weight changes during dietary intervention and colonic inflammation were similar among the experimental groups. Although some studies have reported that feeding with either WD or SCFA alters the weight in obese mice [55], some other studies have suggested that an overweight phenotype does not occur in WD-fed mice [27, 28]. It was postulated that this discrepancy in diet-induced metabolic or body weight changes could be associated with additional experimental factors, including the species or dietary patterns (composition, fat source, micronutrients) [27]. Accordingly, it is possible that westernized diet or SCFA may have caused the rats to feel early satiated and thereby weight gain was prevented [27]. Moreover, SCFA was shown to stimulate the release of the anorectic gut hormones, glucagon-like peptide-1 and peptide YY from human colonic cells [56]. SCFAs are either utilized in the colonocytes, or absorbed into the portal vein or directly into the inferior vena cava to exert their systemic functions, including their roles in metabolic processes, appetite regulation and energy homeostasis [57, 58]. When taken together, our findings suggest that metabolic disturbances that have occurred due to the coexistence of colitis and WD were reversed by SCFA consumption without altering the calorie intake or weight of the animals.

In parallel with the clinical studies showing that portal and mesenteric blood flow are increased in patients with active IBD as compared to controls [59], DSS-treated mice also revealed that the colonic blood flow was increased and vascular resistance was decreased due to colitis [60]. Our current findings also demonstrated that acute oxidative damage of the colon was accompanied by an exaggerated blood flow to the inflamed colon. Furthermore, in the colitis-induced rats that were previously fed with WD or fed afterwards with SCFA, blood flow was depressed in parallel with the alleviation of inflammation. Although experimental studies have reported that SCFAs increase colonic mucosal blood flow [61], current data suggest that the relative reduction in the serosal blood flow by SCFA is secondary to its anti-inflammatory effect. In contrary to several studies reporting that consumption of the WD or high-fat diet aggravates the severity of DSS-induced colitis [7, 62], our findings showed that prior feeding with WD for 3 weeks protected the colon from the injurious effects of DSS, and prevented the increases in lipid peroxidation, ROS generation, neutrophil infiltration and down-regulated the pro-inflammatory cytokine expressions in the colon. Similarly, adding SCFA to normal diet during the inflammatory process decreased the colonic expression of the pro-inflammatory cytokines (IL-6, TNF-α) and increased IL-10 expression, but had no effect on colitis-induced oxidative damage. Despite publications showing that SCFAs induce ROS production, divergent anti-inflammatory and pro-inflammatory effects of SCFAs were also reported [63, 64]. Prolonged intake of the obesogenic diet was reported to impair the gut barrier and cause colonic inflammation [65], while high-fat diet feeding for even 2 weeks has resulted in hepato-intestinal inflammatory state with alterations in bacterial populations [66]. When taken with the aforementioned studies, our current findings suggest that the inflammatory state of the colon induced by consumption of WD could be acting as a preconditioning mechanism and protecting the colonic tissue against the future insults. On the other hand, bowel injury when preceded by WD-induced inflammation has increased neutrophil recruitment to the liver, showing the pro-inflammatory effect of WD diet plus colitis on hepatic tissue. Thus, our findings suggest that feeding with SCFA exerted an anti-inflammatory effect on the colonic tissue, while reduced ROS generation along with enhanced hepatic GSH content indicate an additional antioxidant effect of SCFA on the liver.

The beneficial effects of SCFA in preventing metabolic syndrome and improving the lipid profile were attributed to its regulatory effect on PPAR-γ [67], advocating the use of SCFA as low-cost PPAR-γ modulators [67]. It has been reported that stimulation of PPAR-γ, which is expressed at high levels in the colonic epithelium, decreases the production of pro-inflammatory cytokines by regulating the activity of NF-κB pathway [68, 69]. In accordance with that, ligands for PPAR-γ were shown to reduce colonic inflammation and the levels of pro-inflammatory cytokines in mice with DSS-induced colitis [1, 70-72]. Our results revealed that increased colonic expressions of IL-6 and TNF-α in colitis-induced rats were accompanied by elevated nuclear PPAR-γ2 expressions in the inflamed colonic tissues, suggesting the upregulation of nuclear PPAR-γ2 by the inflammatory insult. Although feeding with SCFA during the course of colonic inflammation depressed the expressions of IL-6 and TNF-α and elevated the expression of IL-10, PPAR-γ1 and PPAR-γ2 expressions were not affected by the presence of SCFA in the diet. On the other hand, alleviated oxidative injury along with depressed expressions of the pro-inflammatory cytokines and elevated IL-10 expression in WD-fed rats was accompanied by the suppression of the nuclear PPAR-γ2 expression. It was shown that PPARs are continuously transported between nucleus and cytoplasm, and the pathways controlling the nuclear–cytoplasmic transport of PPARs could be also involved in the actions of PPARs [73]. Thus, our results suggest that the protective effect of WD against the occurrence of colonic inflammation involves its modulatory impact on nuclear PPAR-γ2 signaling. However, despite that short-term feeding with SCFA inhibited pro-inflammatory cytokine production during the inflammatory challenge, elevated PPAR-γ2 expression was not reduced and oxidative colonic injury was not alleviated. It was previously shown that the anti-inflammatory effect of the PPAR-γ agonists was greater as a preventive treatment than their effect when given after the induction of colitis [70]. Taken together, it can be suggested that both the length and the timing of diet change, either by shifting to WD or by adding SCFA, are critical in aggravating or alleviating the severity of colonic inflammation and involves the modulation of PPAR-γ signaling.

Since diets high in fats were shown to induce colonic oxidative stress, consumption of WD is likely to induce the upregulation of anti-inflammatory signals to combat against an additional oxidative injury. In accordance, our results showed that acute colonic inflammation is alleviated when it is preceded by the consumption of WD, suggesting a preconditioning impact of the WD via the modulation of PPAR-γ activity. Although the beneficial impact of SCFA on colonic inflammation and metabolic diseases was previously demonstrated, we evaluated for the first time that being on WD prior to inflammation or adding SCFA to the diet similarly modulates the cytokine responses, but co-intake of WD and SCFA does not have an additive effect in alleviating oxidative injury. However, since WD-induced protection against colonic inflammation was accompanied by a disrupted lipid profile, which was already impaired by the colitis itself, SCFAs with their beneficial effects on lipid metabolism would be specifically more advantageous in IBD patients comorbid with metabolic syndrome.

Author Contributions: Conceptualization, M.S., B.Y. and A.Y.; methodology, M.S., Ç.Ç.Ö., A.M., E.C.G., E.Y.A., M.Ö., M.T., S.Ş., M.Y., Ö.Ç., B.Ç.Y., A.Y.; writing—original draft preparation, M.S., B.Y. and A.Y.; writing—review and editing, B.Y. and A.Y. All authors have read and agreed to the published version of the manuscript.

Acknowledgments: The authors are grateful to Professor Goncagül Haklar and Dr. Ayşe Mine Yılmaz for their guidance in measuring the parameters in serum samples and in PPAR-γ blotting.

Funding: This research was funded by the Scientific and Technological Research Council of Turkey (TUBITAK–programme code: 2209), grant number 1919B011901268.

STATEMENTS AND DECLARATIONS

Ethics Approval: The study was approved by the Marmara University Animal Care and Use Committee (protocol code 97.2018.mar and date of 5 November 2018) and conducted according to the guidelines of the New York Academy of Science and the ARRIVE guidelines.

Consent for Publication: All authors have read and approved the submission.

Conflicts of Interest: The authors declare no conflict of interest.

1. Wang L, Xie H, Xu L, Liao Q, Wan S, Yu Z, et al. rSj16 protects against DSS-induced colitis by inhibiting the PPAR-α signaling pathway. Theranostics 2017; 7:3446-3446.

2. Knight-Sepulveda K, Kais S, Santaolalla R, Abreu MT. Diet and inflammatory bowel disease. Gastroenterology and Hepatology, 2015.

3. Bernstein CN, Eliakim A, Fedail S, Fried M, Gearry R, Goh KL, et al. World gastroenterology organisation global guidelines inflammatory bowel disease. Journal of Clinical Gastroenterology, 2016.

4. Statovci D, Aguilera M, MacSharry J, Melgar S. The impact of western diet and nutrients on the microbiota and immune response at mucosal interfaces. Frontiers in Immunology, 2017.

5. Hanauer SB. Inflammatory bowel disease: epidemiology, pathogenesis, and therapeutic opportunities. Inflammatory bowel diseases 2006; 12:S3-S9.

6. Amre DK, D'Souza S, Morgan K, Seidman G, Lambrette P, Grimard G, et al. Imbalances in dietary consumption of fatty acids, vegetables, and fruits are associated with risk for crohn's disease in children. American Journal of Gastroenterology 2007.

7. Agus A, Denizot J, Thévenot J, Martinez-Medina M, Massier S, Sauvanet P, et al. Western diet induces a shift in microbiota composition enhancing susceptibility to Adherent-Invasive E. coli infection and intestinal inflammation. Scientific Reports 2016.

8. Lee D, Albenberg L, Compher C, Baldassano R, Piccoli D, Lewis JD, et al. Diet in the pathogenesis and treatment of inflammatory bowel diseases. Gastroenterology 2015; 148:1087-1106.

9. Fengming Y, Jianbing W. Biomarkers of inflammatory bowel disease. Disease markers 2014; 2014.

10. Strober W, Fuss I, Mannon P. The fundamental basis of inflammatory bowel disease. The Journal of clinical investigation 2007; 117:514-521.

11. Kretzmann NA, Fillmann H, Mauriz JL, Marroni CA, Marroni N, González-Gallego J, et al. Effects of glutamine on proinflammatory gene expression and activation of nuclear factor kappa B and signal transducers and activators of transcription in TNBS-induced colitis. Inflammatory bowel diseases 2008; 14:1504-1513.

12. Macfarlane S, Macfarlane GT. Regulation of short-chain fatty acid production. Proceedings of the Nutrition Society 2003.

13. Miller TL, Wolin MJ. Pathways of acetate, propionate, and butyrate formation by the human fecal microbial flora. Applied and Environmental Microbiology 1996.

14. Louis P, Flint HJ. Formation of propionate and butyrate by the human colonic microbiota. Environmental Microbiology, 2017.

15. Cummings JH, Pomare EW, Branch HWJ, Naylor CPE, MacFarlane GT. Short chain fatty acids in human large intestine, portal, hepatic and venous blood. Gut 1987.

16. Bergman EN. Energy contributions of volatile fatty acids from the gastrointestinal tract in various species. Physiological Reviews, 1990.

17. Kripke SA, Fox AD, Berman JM, Settle RG, Rombeau JL. Stimulation of Intestinal Mucosal Growth with Intracolonic Infusion of Short-Chain Fatty Acids. Journal of Parenteral and Enteral Nutrition 1989.

18. Gonzalez A, Krieg R, Massey HD, Carl D, Ghosh S, Gehr TWB, et al. Sodium butyrate ameliorates insulin resistance and renal failure in CKD rats by modulating intestinal permeability and mucin expression. Nephrology, dialysis, transplantation : official publication of the European Dialysis and Transplant Association - European Renal Association 2019.

19. Zhang T, Xia M, Zhan Q, Zhou Q, Lu G, An F. Sodium Butyrate Reduces Organ Injuries in Mice with Severe Acute Pancreatitis Through Inhibiting HMGB1 Expression. Dig Dis Sci 2015; 60:1991-9.

20. Cummings JH, Beatty ER, Kingman SM, Bingham SA, Englyst HN. Digestion and physiological properties of resistant starch in the human large bowel. British Journal of Nutrition 1996.

21. Sivaprakasam S, Gurav A, Paschall AV, Coe GL, Chaudhary K, Cai Y, et al. An essential role of Ffar2 (Gpr43) in dietary fibre-mediated promotion of healthy composition of gut microbiota and suppression of intestinal carcinogenesis. Oncogenesis 2016.

22. den Besten G, Lange K, Havinga R, van Dijk TH, Gerding A, van Eunen K, et al. Gut-derived short-chain fatty acids are vividly assimilated into host carbohydrates and lipids. American Journal of Physiology - Gastrointestinal and Liver Physiology 2013.

23. Mathewson ND, Jenq R, Mathew AV, Koenigsknecht M, Hanash A, Toubai T, et al. Gut microbiome-derived metabolites modulate intestinal epithelial cell damage and mitigate graft-versus-host disease. Nature Immunology 2016.

24. Asarat M, Vasiljevic T, Apostolopoulos V, Donkor O. Short-Chain Fatty Acids Regulate Secretion of IL-8 from Human Intestinal Epithelial Cell Lines in vitro. Immunological Investigations 2015.

25. Kim MH, Kang SG, Park JH, Yanagisawa M, Kim CH. Short-chain fatty acids activate GPR41 and GPR43 on intestinal epithelial cells to promote inflammatory responses in mice. Gastroenterology 2013.

26. Canfora EE, Jocken JW, Blaak EE. Short-chain fatty acids in control of body weight and insulin sensitivity. Nature Reviews Endocrinology, 2015.

27. Hintze KJ, Benninghoff AD, Cho CE, Eward R. Modeling the western diet for preclinical investigations. Advances in Nutrition, 2018.

28. Monsanto SP, Hintze KJ, Ward RE, Larson DP, Lefevre M, Benninghoff AD. The new total Western diet for rodents does not induce an overweight phenotype or alter parameters of metabolic syndrome in mice. Nutrition Research 2016.

29. Chassaing B, Aitken JD, Malleshappa M, Vijay-Kumar M. Dextran sulfate sodium (DSS)-induced colitis in mice. Current Protocols in Immunology 2014.

30. Kiesler P, Fuss IJ, Strober W. Experimental models of inflammatory bowel diseases. Medecine et Hygiene, 2001.

31. Vicario M, Crespí M, Franch À, Amat C, Pelegrí C, Moretó M. Induction of colitis in young rats by dextran sulfate sodium. Digestive diseases and sciences 2005; 50:143-150.

32. Ichikawa H, Shineha R, Satomi S, Sakata T. Gastric or rectal instillation of short-chain fatty acids stimulates epithelial cell proliferation of small and large intestine in rats. Digestive Diseases and Sciences 2002.

33. Vieira ELM, Leonel AJ, Sad AP, Beltrão NRM, Costa TF, Ferreira TMR, et al. Oral administration of sodium butyrate attenuates inflammation and mucosal lesion in experimental acute ulcerative colitis. Journal of Nutritional Biochemistry 2012.

34. Zani A, Cordischi L, Cananzi M, De Coppi P, Smith VV, Eaton S, et al. Assessment of a neonatal rat model of necrotizing enterocolitis. European Journal of Pediatric Surgery 2008.

35. Pamukcu O, Kumral ZNO, Ercan F, Yegen BÇ, Ertem D. Anti-inflammatory effect of obestatin and ghrelin in dextran sulfate sodium-induced colitis in rats. Journal of Pediatric Gastroenterology and Nutrition 2013.

36. Bradley PP, Priebat DA, Christensen RD, Rothstein G. Measurement of cutaneous inflammation: estimation of neutrophil content with an enzyme marker. J Invest Dermatol 1982; 78:206-9.

37. Beutler E. Glutathione in Red Cell Metabolism : A manual of Biochemical methods, 1975.

38. Aykac G, Uysal M, Yalcin AS, Kocak-Toker N, Sivas A, Oz H. The effect of chronic ethanol ingestion on hepatic lipid peroxide, glutathione, glutathione peroxidase and glutathione transferase in rats. Toxicology 1985; 36:71-6.

39. Deshpande SS. Principles and applications of luminescence spectroscopy. Critical Reviews in Food Science and Nutrition 2001.

40. Arabacı Tamer S, Yıldırım A, Arabacı Ş, Çiftçi S, Akın S, Sarı E, et al. Treatment with estrogen receptor agonist ERβ improves torsion-induced oxidative testis injury in rats. Life Sciences 2019.

41. Charan J, Kantharia ND. How to calculate sample size in animal studies? J Pharmacol Pharmacother 2013; 4:303-6.

42. Festing MF, Altman DG. Guidelines for the design and statistical analysis of experiments using laboratory animals. ILAR J 2002; 43:244-58.

43. Arifin WN, Zahiruddin WM. Sample Size Calculation in Animal Studies Using Resource Equation Approach. Malays J Med Sci 2017; 24:101-105.

44. Festing MF. On determining sample size in experiments involving laboratory animals. Lab Anim 2018; 52:341-350.

45. Hollander M, Wolfe DA. Nonparametric statistical methods. New York: Wiley, 1999:xiv, 787 p.

46. Krishnan A, Korzenik JR. Inflammatory bowel disease and environmental influences. Gastroenterol Clin North Am 2002; 31:21-39.

47. Persson PG, Ahlbom A, Hellers G. Diet and inflammatory bowel disease: a case-control study. Epidemiology 1992; 3:47-52.

48. Kang EA, Han K, Chun J, Soh H, Park S, Im JP, et al. Increased Risk of Diabetes in Inflammatory Bowel Disease Patients: A Nationwide Population-based Study in Korea. J Clin Med 2019; 8.

49. Soh H, Chun J, Han K, Park S, Kang EA, Im JP, et al. P797 Crohn’s disease and ulcerative colitis was associated with different lipid profile disorders: a nationwide population-based study. Journal of Crohn's and Colitis 2019; 13:S521-S521.

50. Gabbia D, Roverso M, Guido M, Sacchi D, Scaffidi M, Carrara M, et al. Western diet-induced metabolic alterations affect circulating markers of liver function before the development of steatosis. Nutrients 2019.

51. Vahidinia A, Komaki H, Rahbani M, Darabi M, Mahjub H. Effects of dietary garlic supplements on serum lipid profles, LDL oxidation and weight gain in Western diet-fed rats. Progress in Nutrition 2017.

52. Fushimi T, Suruga K, Oshima Y, Fukiharu M, Tsukamoto Y, Goda T. Dietary acetic acid reduces serum cholesterol and triacylglycerols in rats fed a cholesterol-rich diet. Br J Nutr 2006; 95:916-24.

53. Lu YY, Fan C, Li P, Lu YY, Chang X, Qi K. Short Chain Fatty Acids Prevent High-fat-diet-induced Obesity in Mice by Regulating G Protein-coupled Receptors and Gut Microbiota. Scientific Reports 2016; 6:37589-37589.

54. Kondo T, Kishi M, Fushimi T, Ugajin S, Kaga T. Vinegar intake reduces body weight, body fat mass, and serum triglyceride levels in obese Japanese subjects. Biosci Biotechnol Biochem 2009; 73:1837-43.

55. Lu Y, Fan C, Liang A, Fan X, Wang R, Li P, et al. Effects of SCFA on the DNA methylation pattern of adiponectin and resistin in high-fat-diet-induced obese male mice. British Journal of Nutrition 2018.

56. Chambers ES, Viardot A, Psichas A, Morrison DJ, Murphy KG, Zac-Varghese SE, et al. Effects of targeted delivery of propionate to the human colon on appetite regulation, body weight maintenance and adiposity in overweight adults. Gut 2015; 64:1744-54.

57. Haghikia A, Jörg S, Duscha A, Berg J, Manzel A, Waschbisch A, et al. Dietary Fatty Acids Directly Impact Central Nervous System Autoimmunity via the Small Intestine. Immunity 2015.

58. Byrne CS, Chambers ES, Morrison DJ, Frost G. The role of short chain fatty acids in appetite regulation and energy homeostasis. Int J Obes (Lond) 2015; 39:1331-8.

59. Maconi G, Imbesi V, Bianchi Porro G. Doppler ultrasound measurement of intestinal blood flow in inflammatory bowel disease. Scand J Gastroenterol 1996; 31:590-3.

60. Garrelds IM, Heiligers JP, Van Meeteren ME, Duncker DJ, Saxena PR, Meijssen MA, et al. Intestinal blood flow in murine colitis induced with dextran sulfate sodium. Dig Dis Sci 2002; 47:2231-6.

61. Scheppach W. Effects of short chain fatty acids on gut morphology and function. Gut 1994; 35:S35-8.

62. Lee JC, Lee HY, Kim TK, Kim MS, Park YM, Kim J, et al. Obesogenic diet-induced gut barrier dysfunction and pathobiont expansion aggravate experimental colitis. PLoS ONE 2017.

63. Fang W, Xue H, Chen X, Chen K, Ling W. Supplementation with sodium butyrate modulates the composition of the gut microbiota and ameliorates high-fat diet-induced obesity in mice. Journal of Nutrition 2019.

64. Vinolo MAR, Rodrigues HG, Nachbar RT, Curi R. Regulation of inflammation by short chain fatty acids. Nutrients, 2011.

65. Lam YY, Ha CW, Campbell CR, Mitchell AJ, Dinudom A, Oscarsson J, et al. Increased gut permeability and microbiota change associate with mesenteric fat inflammation and metabolic dysfunction in diet-induced obese mice. PLoS One 2012; 7:e34233.

66. Yildirim A, Arabaci Tamer S, Sahin D, Bagriacik F, Kahraman MM, Onur ND, et al. The effects of antibiotics and melatonin on hepato-intestinal inflammation and gut microbial dysbiosis induced by a short-term high-fat diet consumption in rats. Br J Nutr 2019; 122:841-855.

67. den Besten G, Bleeker A, Gerding A, van Eunen K, Havinga R, van Dijk TH, et al. Short-Chain Fatty Acids Protect Against High-Fat Diet-Induced Obesity via a PPARgamma-Dependent Switch From Lipogenesis to Fat Oxidation. Diabetes 2015; 64:2398-408.

68. Sanchez-Hidalgo M, Martin AR, Villegas I, Alarcon De La Lastra C. Rosiglitazone, an agonist of peroxisome proliferator-activated receptor gamma, reduces chronic colonic inflammation in rats. Biochem Pharmacol 2005; 69:1733-44.

69. Torres J, Danese S, Colombel JF. New therapeutic avenues in ulcerative colitis: thinking out of the box. Gut 2013; 62:1642-52.

70. Bento AF, Marcon R, Dutra RC, Claudino RF, Cola M, Leite DFP, et al. β-caryophyllene inhibits dextran sulfate sodium-induced colitis in mice through CB2 receptor activation and PPARγ pathway. American Journal of Pathology 2011.

71. Shen P, Zhang Z, He Y, Gu C, Zhu K, Li S, et al. Magnolol treatment attenuates dextran sulphate sodium-induced murine experimental colitis by regulating inflammation and mucosal damage. Life Sciences 2018.

72. Su CG, Wen X, Bailey ST, Jiang W, Rangwala SM, Keilbaugh SA, et al. A novel therapy for colitis utilizing PPAR-gamma ligands to inhibit the epithelial inflammatory response. J Clin Invest 1999; 104:383-9.

73. Umemoto T, Fujiki Y. Ligand-dependent nucleo-cytoplasmic shuttling of peroxisome proliferator-activated receptors, PPARα and PPARγ. Genes to Cells 2012.

The Effect of Short-chain Fatty Acid Supplementation on Acute Colonic Inflammation in Rats Fed With Western Diet: Role of the Peroxisome Proliferator-activated Receptor-gamma

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Abstract

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1. Introduction

2. Material And Methods

3. Results

4. Discussion

5. Conclusion

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