Clostridium Tyrobutyricum Protect Intestinal Barrier Function from LPS-Induced Apoptosis via P38/JNK Signaling Pathway in IPEC-J2 Cells

The intestinal epithelial barrier, which is formed by intestinal mucosa, separates the internal milieu from the external environment and is the first line of defense against bacterial and toxin invasion of host tissues [1, 2]. At early weaning, the intestinal barrier function of pigs is vulnerable to damage. The epithelial layer impairment results in poor nutrient digestion and absorption, reduced transmucosal resistance, imbalanced secretion and absorption of water and electrolytes, and increased enteric pathogen infection. Disruption of the intestinal barrier disturbs immune homeostasis, increases inflammation in the intestine, and is associated with intestinal diseases such as diarrhea [3-5]. Therefore, maintenance of intestinal barrier function plays a pivotal role in optimum gastrointestinal health.

Probiotics, defined as live microorganisms, can maintain epithelial integrity and barrier function, stimulate the cellular repair mechanism, and reestablish well-balanced indigenous intestinal and respiratory microbial communities [6]. Probiotics such as Lactobacillus and Enterococcus faecium help to protect the intestinal epithelium after infection with E. coli or exposure to lipopolysaccharide (LPS) [7, 8]. LPS, produced by Gram-negative bacteria, plays important roles in the integrity of the outer membrane permeability barrier and participates extensively in host-pathogen interplay [9]. LPS interacts with and activates Tolllike receptors, which themselves activate mitogen-activated protein kinases (MAPK) [10]. The MAPK family can be classified into extracellular signal-regulated kinase (ERK), c-Jun N-terminal kinase (JNK), and p38. Signaling by these pathways govern fundamental cellular responses, such as cell proliferation, differentiation, inflammation, and apoptosis [11-12].

Clostridium tyrobutyricum(C.t), a Gram-positive anaerobic bacterium, can efficiently produce butyric acid and is a promising probiotic to protect intestinal epithelial function. Therefore, in this study, we investigated whether C.t could protect porcine intestinal epithelial barrier function, hypothesizing that its benefit might be associated with the MAPK signaling pathway in IPEC-J2 cells.

Intestinal porcine epithelial cells (IPEC-J2) were obtained from the Institute of Subtropical Agriculture (Chinese Academy of Sciences) and cultured in Dulbecco's modified Eagle medium/F12 medium (Biological Industries, Kibbutz Beit Haemek, Israel) supplemented with 10% fetal bovine serum (Gemini, USA), 100 U/mL penicillin, and 100 µg/mL streptomycin (Biological Industries, Kibbutz Beit Haemek, Israel) at 37 °C with 5% CO₂ in a humidified atmosphere. No antibiotics were used during the treatment. C.t was obtained from Ohio State University and cultured anaerobically at 37°C in clostridial growth medium (CGM) [13]. C.t was then collected after centrifugation at 12, 000 rpm for 5 min, washed 3 times, and suspended in phosphate-buffered saline(PBS) at different concentrations (10⁶ to 10¹¹ CFU/mL) for cell treatments.

Cell viability following treatment with different concentrations of C.t (ranging from 10⁶ to 10¹¹ CFU/ mL) for different hours (2, 4, 6, 8, and 10 h) was measured using the Cell Counting Kit 8 (MedChem Express, Monmouth Junction, NJ, USA). Cell cytotoxicity following the treatments with different concentrations of C.t (ranging from 10⁸ to 10¹¹ CFU/mL) for 8 h was measured using a lactate dehydrogenase (LDH) detection kit (Beyotime, Shanghai, China) according to the manufacturer's instructions. Briefly, IPEC-J2 cells were seeded in 96-well plates at a density of 1 × 10⁴ cells/well, cultured overnight to allow cell attachment, and then pretreated with C.t after reaching 70-80% confluence. To evaluate cell viability, 10 µL of the Cell Counting Kit 8 assay solution was added to each treated cell well and the cells were further incubated for 1 to 2 h. Subsequently, the optical density was measured using SpectraMax M5 (Molecular Devices, California, USA) at a 450 nm wavelength‚and the percentage of living cells was calculated as described by Mosmann [14]. As for cell cytotoxicity, 150 µL LDH release solution was added to each cell well and incubated for 1 h. Next, 120 µL cell-free culture supernatants were collected into new 96-well plates after centrifugation at 1, 200 rpm for 5 min. Finally, 60 µL LDH working assay solution was added to each cell well and the optical density was measured using the SpectraMaxM5 at a 490 nm wavelength with a reference wavelength of 620 nm [15].

IPEC-J2 cells were seeded onto polycarbonate membrane filters (0.4-µm pore size, 1.12 cm² growth area) inside Transwell® cell culture chambers (Corning Costar, Cambridge, MA, USA) at a density of 1 × 10⁵ cells/cm². The culture medium was changed every day and trans-epithelium electrical resistance (TEER) values were measured every other day using the Millicell Electrical Resistance System (Millipore ERS-2, Massachusetts, USA). When a monolayer of cells was considered to be completely differentiated as previously described [16], cells were treated with PBS, LPS (LPS; 1 µg/mL; LPS 055:B55, Sigma, Massachusetts, USA), C.t, LPS + C.t (cells were treated with 1 µg/mL LPS, followed by C.t), or C.t + LPS (cells treated with C.t, followed by 1 µg/mL LPS), and the TEER values were measured every 4 h.

Immediately after the last TEER value measurement, 100 µL of 1 mg/mL 4-kDa fluorescein isothiocyanate-dextran (FD4; Sigma-Aldrich, St. Louis, MO, USA) was added into the upper compartments of the Transwells. The Transwells were cultured at 37 °C for 30 min. Then, 100 µL of medium of each well from the lower compartments of the Transwells was added into wells of a black 96-well plate and the fluorescence intensity was detected using a SpectraMax M5 at an excitation wavelength of 480 nm and emission wavelength of 520 nm [17].

Apoptotic IPEC-J2 cells were detected using an Annexin V-FITC/PI kit (R&D Systems, Minnesota, USA) according to the manufacturer's instructions. Cells at a density of 1 × 10⁶ cells/well were seeded in 6-well plates overnight. After the treatments were finished, 500 µL of 1 × Binding Buffer, 5 µL Annexin V-FITC, and 5 µL propidium iodide (PI) were added into each well after two washes with PBS and the plates were incubated for 15 min at room temperature in the dark. Fluorescence images were taken by a TE2000 fluorescence microscope (Nikon, Tokyo‚Japan). To determine apoptotic cells by flow cytometry cells at a density of 1 × 10⁶ cells/well were seeded in 6-well plates overnight. The medium and cells were digested by pancreatin without ethylenediaminetetraacetic acid (EDTA) (Beyotime), collected into 10-mL centrifugal tubes, and centrifuged for 5 min at 1, 000 rpm. Cells were then suspended in 100 µLl × Binding Buffer at a concentration of 1 × 10⁵ cells/mL in 10-mL centrifugal tubes. Next, 5 µL FITC Annexin V and 5 µL PI were added into the tubes and they were incubated for 15 min at room temperature in the dark. Finally, 400 µL 1 × Binding Buffer was added to each tube and cell apoptosis was analyzed by flow cytometry (FACSCalibur, B&D, Minnesota, USA) within 1 h.

Cells cultured in Transwells and 6-cell culture plates were first digested with 0.25% trypsin/EDTA (Biological Industries, Kibbutz Beit Haemek, Israel) into centrifuge tubes. Total cell protein was extracted using the Whole Cell Lysis Assay Kit (KeyGEN BioTech, Beijing, China) after two PBS washes. The protein concentration was determined using the BCA Assay Kit (KeyGEN BioTech). Approximately 30 µg of protein was boiled in 5 × loading buffer (Sangon Biotech, Shanghai, China), separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis, and then transferred to polyvinylidene fluoride membranes (Millipore, Massachusetts, USA). The membranes were blocked with 5% non-fat milk for 1 h at room temperature and then incubated in the corresponding primary antibodies at 4 °C overnight. After being washed with TBST 3 times for 10 mineach, the membranes were incubated with secondary antibodies for 1 h at room temperature. The membranes were then washed with TBST as before. The immunoreactive bands were visualized with ECL luminescence reagent (KeyGEN BioTech) using a ChemiScope Touch (Clinx, Shanghai, China). The band intensities were determined using ImageJ software. Primary antibodies ZO-1, Claudin-1, and Occluding were purchased from Proteintech (Wuhan‚China). ERK1/2, phospho-ERKl/2, p-38, phospho-p38, JNK, phospho-JNK, and HRP-conjugated anti-rabbit IgG were obtained from Cell Signaling Technology (Massachusetts, USA), β-actin was purchased from Abeam (Massachusetts, USA), and HRP-conjugated anti-mouse IgG was obtained from Beyotime.

Total RNA was extracted using TRIzol reagent (Invitrogen, Carlsbad, California, USA). RNA quantity and quality were determined using a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, Massachusetts, USA). cDNA was synthesized with the PrimeScript RT reagent kit with gDNA Eraser (TAKARA, Dalian, China) according to the manufacturer's instructions. Briefly, 1 µg total RNA was used to erase gDNA at 42°C for 2 min. The reverse transcription was conducted at 37 °C for 15 min and 85°C for 5 s. Real-time PCR was performed on a CFX96TM Real-Time System (Bio-Rad, Hercules, CA, USA) in triplicate, in a total volume of 25 µL consisting of 12.5 µL SYBR Premix EX Taq (TAKARA), 0.5 µL of each primer (10 µM), 2 µL of cDNA template, and 9.5 µL double-distilled water. The PCR cycle conditions were 95 °C for 30 s, followed by 40 cycles of 95 °C for 5 s and 60 °C for 30 s. Melting curve analysis was used to confirm the specificity and reliability of the PCR products, β-actin was used as a house-keeping gene to normalize target gene levels. The relative mRNA expression was calculated using the 2^-∆∆Ct method [18]. Primers used in this study were designed with Primer 5.0 (Table 1) and synthesized in Tsingke (Beijing, China).

Statistical analysis was performed with one-way analysis of variance followed by a Duncan multiple range test with SPSS 17.0 (SPSS, Chicago, IL, USA). All data are expressed as the mean ± standard error of the mean (SEM). Differences were considered significant at P < 0.05.

IPEC-J2 cells were treated with C.t at different final concentrations (ranging from 10⁶ to 10¹¹ CFU/mL) for 2, 4, 6, 8, and 10 h. As shown in Fig. 1A, C.t treatment enhanced the growth of IPEC-J2 cells in a dose- and time-dependent manner. For instance, cell viability was increased significantly at 8 h (P < 0.001). When cells were treated with high concentrations (ranging from 10⁸ to 10¹¹ CFU/mL), cell viability was increased significantly compared with other concentrations (P < 0.05) (Fig. 1A). Cell cytotoxicity was measured after cells were treated with C.t (ranging from 10⁸ to 10¹¹ CFU/mL) for 8 h, with the results showing that LDH release was decreased when cells were treated with 10⁸ and 10⁹ CFU/mL C.t compared with the control (P < 0.01; Fig. 1B). Therefore, IPEC-J2 cells treated with 10⁸ CFU/mL C.t for 8 h were used in the subsequent experiments.

An intestinal barrier model was used to investigate whether C.t could inhibit the in vitro LPS-induced increase in intestinal permeability. When the TEER value reached 1524.43 ± 15.02 Ω·cm², the monolayer was considered to be completely differentiated (Fig. 2A). Cells were treated with PBS (the control), LPS, C.t, LPS + C.t, or C.t + LPS (Fig. 2B), and TEER measurements were conducted every 4 hours. The TEER value reduced to the lowest (∼40 Ω·cm²) in 20 h with LPS treatment, indicating that the intestinal integrity was greatly disturbed. No significant differences were detected between the control and the C.t treatment, implying that C.t positively maintained the TEER values of IPEC-J2 cells with time. As for the LPS + C.t treatment, the TEER value decreased to 217 Ω·cm² in the first 12 h when cells were pretreated with LPS. However, the TEER value was maintained at about 223 Ω·cm² after the cells were treated with C.t in the next 8 h. For the cells treated with C.t + LPS, the TEER value was the same as the control when the cells were pretreated with C.t in the first 8 h, whereas the TEER value reached the minimum after LPS was added in the next 12 h (Fig. 2C).

After the TEER values were measured, FD4 was added into the upper compartments of the Transwells to ensure that C.t could suppress the LPS-induced increase in intestinal permeability. The concentration of FD4 was markedly increased by LPS treatment compared with the control and C.t treatment groups (P < 0.01) and no differences were observed between the control and C.t-treated cells. In addition, the concentrations of FD4 were decreased by the LPS + C.t and C.t + LPS treatments compared with LPS alone (P < 0.01; Fig. 2D).

The expressions of tight junction (TJ) proteins (ZO-1, Claudin-1, and Occludin) were also measured in this study. After treatment with LPS or C.t for 20 h, the expressions of TJ proteins were significantly decreased in cells treated with LPS (P < 0.05), whereas C.t increased the expressions of all three proteins (P < 0.01). In addition, we noted that C.t inhibited the LPS-induced decrease in TJ protein expressions (Fig. 2E–G).

TJ protein expressions were measured to further investigate the protective effect of C.t on the intestinal barrier. Compared with the control, cells treated with LPS alone for 12 h showed decreased expressions of TJ proteins (P < 0.01; Fig. 3), which was consistent with the TEER, FD4, and TJ protein expression results obtained previously (Fig. 2). No differences could be observed between the control and 8-hC.t treatment groups in the expressions of ZO-1 and Claudin-1, but Occludin expression was much higher in the C.t group than in the control (P < 0.01). Cells treated with LPS + C.t showed increased expression of these three proteins compared with the LPS-alone group, whereas the expressions of TJ proteins werenot different between the C.t + LPS and LPS treatments.

The bright-field images of cells pretreated with LPS showed nuclear and cytoplasmic condensation and some cellular fragments, which are typical characteristics of apoptosis. However, there was no characteristic difference between the control and C.t treatment groups (Fig. 4A). An annexin V-FITC/PI assay was conducted to further confirm whether C.t could attenuate LPS-induced apoptosis. Fluorescence images of FITC-/PI-stained apoptotic IPEC-J2 cells showed that LPS considerably stimulated cell apoptosis. No differences were observed between the control and C.t treatments. Both LPS + C.t and C.t + LPS treatments partially inhibited the LPS-induced apoptosis (Fig. 4B).

The results of flow cytometry quantitatively verified the above results. LPS significantly increased (P < 0.05) the ratio of apoptotic cells, whereas C.t efficiently inhibited apoptosis induced by LPS (Fig. 4C). The flow cytometry results showed that 57.47% cells were alive when the cells were treated LPS alone, which was lower than the percentages in the control and C.t groups (94.4% live cells and 96.8% live cells, respectively; P < 0.001; Fig. 4D and E).

We next analyzed the relative mRNA expression of several apoptotic markers, including caspase-3, caspase-8, Bax, and Bcl-2. As for the expression of the anti-apoptotic gene Bcl-2, the results showed that the relative Bcl-2 expression was significantly decreased with LPS compared with that in the control and C.t treatment groups (P < 0.05). In particular, C.t significantly increased the Bcl-2 expression compared with the control (P < 0.05), signifying its anti-apoptotic effects. Thus, for the LPS + C.t and C.t + LPS treatments, Bcl-2 expressions were higher than with LPS, although the differences were not significant (Fig. 5A). The relative expressions of the pro-apoptotic gene Bax and the apoptotic genes caspase-3/-8 were significantly increased in cells pretreated with LPS compared with the control and C.t treatment groups (P < 0.01). C.t reduced the expression of these apoptosis-related genes (Fig. 5B-D).

The apoptosis inhibitor Z-VAD(OMe)-FMK, a pan-caspase inhibitor, was used in this study to investigate whether C.t could protect intestinal barrier function from LPS-induced apoptosis. The relative expressions of the apoptotic genes caspase-3/-8 were significantly decreased in LPS-treated cells after an apoptosis inhibitor was added (P < 0.05). However, no significant differences were observed in the relative expressions of Bax and Bcl-2 between the LPS and apoptosis inhibitor + LPS treatments. These results indicated that an apoptosis inhibitor could partly prevent LPS-induced apoptosis (Fig. 6A). The expressions of TJ proteins were increased dramatically after the addition of the apoptosis inhibitor to LPS-treated cells (P < 0.05), which were similar to the results of C.t-treated cells (Fig. 6B and C).

To elucidate the mechanisms underlying the effects of C.t on LPS-induced apoptosis, we examined the effects of C.t on the MAPK signaling pathway. Phosphorylation of p38 (p-p38) and JNK (p-JNK) was observed after LPS treatment of cells. However, C.t inhibited the LPS-induced phosphorylation of p-p38 and p-JNK (Fig. 7A–D). We then measured the relative mRNA expressions of downstream genes in this pathway. The mRNA levels of AP-1, ATF-2, ELK-1, and p53 were increased (P< 0.05) and that of Stat3 was decreased (P < 0.001) in cells treated with LPS compared with the control, whereas C.t inhibited these changes (Fig. 7E–I).

Pharmacological inhibitors of p38 (SB203580) and JNK (SP600125) were used to investigate the underlying mechanisms of the protective effects of C.t on LPS-induced apoptosis in IPEC-J2 cells. The concentrations of p-p38 and p-JNK were suppressed when cells were treated with inhibitors or inhibitors + LPS (Fig. 8A and D). The mRNA levels of the anti-apoptotic gene Bcl-2 were increased in cells treated with inhibitors or with inhibitors + LPS compared with cells treated with LPS alone (P < 0.05). In addition, relative mRNA expressions of apoptosis-related genes, including Bax, caspase-3, and caspase-8, were significantly decreased when inhibitors were added to the cells (P < 0.01; Fig. 8B and E). TJ proteins levels were also determined to verify whether C.t could protect intestinal barrier function from LPS-induced apoptosis via the p38/JNK signaling pathway. The results showed that expressions of TJ proteins were increased when p38/JNK signaling pathways were switched off (P < 0.05; Fig. 8C and F).

C.t, a Gram-positive and strictly anaerobic bacterium, has received much attention from the dairy and chemical synthesis industries because of its efficiency in butyric acid production [19, 20]. However, the role of C.t in animal models or mammalian cells has not yet been reported. In this work, the biological function of C.t was investigated in vitro, with the results showing that C.t could protect the intestinal barrier from LPS-induced apoptosis in IPEC-J2 cells.

The intestinal epithelium, which represents the largest interface in the body, plays several important roles in the absorption of nutrients and prevention of water and electrolyte loss [21, 22]. In particular, the intestinal barrier can separate the internal milieu from the external environment via cell-cell adhesion and intercellular junctions that direct morphogenesis and maintain tissue integrity [2]. Intestinal permeability is controlled by epithelial adherens junctions and TJs, with the increased intestinal permeability contributing to the severity of some diseases, such as inflammatory bowel disease and irritable bowel syndrome [21]. In the present study, an in vitro-simulated intestinal barrier was established by culturing IPEC-J2 cells in Transwells. LPS has a negative effect on the TEER values and TJ protein contents of IPEC-J2 cells, ultimately resulting in increased permeability and intestinal barrier dysfunction [23, 24]. Many reports have also revealed that the TEER values and TJ protein expressions markedly decreased after cells were treated with LPS for 12 or 24 h [7, 25]. In our study, we successfully constructed a model of LPS-induced increased permeability. The TEER values decreased to their lowest level and the expression of these TJ proteins was completely reduced after the IPEC-J2 cells were treated with LPS for 20 h. In contrast, C.t stably maintained the TEER values and increased the expression of the TJ protein sat 20 h. The decrease in the TEER values and TJ protein expression were also inhibited by C.t. Our further research on the expression of TJ proteins in cells treated with LPS for only 12 h or with C.t for 8 h also verified the protective function of C.t. These results indicate that C.t protected the intestinal barrier from the LPS-induced increase in intestinal permeability.

Apoptosis, which is a type of programmed cell death involving the activation of apoptotic caspases, can be initiated by a variety of stimuli arising from receptors on the cell surface or from the detection of events within the cell [26]. Tumor necrosis factor (TNF) or LPS stimulation increase apoptosis and consequent cell shedding which are associated with barrier loss [27]. We noticed in the present study that, after treatment with LPS, cells showed nuclear and cytoplasmic condensation, typical characteristics of apoptosis. LPS, which is a major component in the outer monolayer of most Gram-negative bacteria, can induce apoptosis and disturb intestinal integrity in IPEC-J2 cells [28]. Thus, in the present study, we hypothesized that C.t could inhibit LPS-induced apoptosis and ultimately protect the intestinal barrier. We found that LPS induced apoptosis at 12 h and that C.t efficiently inhibited this phenomenon. The relative mRNA expression levels of apoptosis-related genes, including Bcl-2, Bax, and caspase-3/-8, were also measured in this study; their levels were consistent with the results of the Annexin V-FITC/PI assay and flow cytometry Moreover, intestinal barrier function was improved when LPS-induced apoptosis was prevented by a pharmacological apoptotic inhibitor. Our results qualitatively and quantitatively indicated that C.t may protect intestinal barrier function by markedly suppressing LPS-induced apoptosis in IPEC-J2 cells.

MAPKs play essential roles in some physiological cell processes, including inflammation, stress, growth, differentiation, and death. ERK1/2 are mostly activated by growth factors, such as epidermal growth factor and nerve growth factor, which play central roles in the control of cell proliferation [29, 30]. While p38 and JNK, known as stress-activated protein kinase, are strongly activated by various cellular stresses, such as heat shock and oxidative stress, TNF-α and LPS, which play roles in cell proliferation and survival, are associated with the induction of apoptosis by cellular stresses [30]. In the present study, p38 and JNK phosphorylation occurred after cells were treated with LPS for 12 h. The major functional role of phosphorylated p38 and JNK lies in mediating the downstream substrates involved in the cell cycle and apoptosis [29]. Our results also suggest that the relative mRNA expressions of downstream genes of the p38 and JNK signaling pathways, including AP-1, ATF-2, ELK-1, and p53, were increased and that Stat3 expression was dramatically decreased after cells were treated with LPS, which may activate caspases and induce apoptosis [31, 32]. Pharmacological inhibitors were used to switch off the activation of p38/JNK signaling pathways; we found that these inhibitors could restrain LPS-induced apoptosis and thus improve intestinal barrier function. Therefore, we speculated that C.t inhibited LPS-induced apoptosis by depressing the p38/JNK signaling pathway (Fig. 9). However, the molecular basis of how C.t suppresses the activity of the p38/JNK signaling pathway remains to be investigated.

Our findings suggest that C.t could protect against the LPS-induced increase in intestinal permeability and inhibit LPS-induced intestinal apoptosis via the p38/JNK signaling pathway in IPEC-J2 cells. These results suggest C.t is a potential probiotic additive for protecting the intestinal barrier and preventing diarrhea in early weaned piglets.

This study was supported by the Project of Ministry of Agriculture (948) (Grant No. 2014-Z27).

We would like to thank Shaojuan Liu from Institute of Subtropical Agriculture, Chinese Academy of Sciences for IPEC-J2 cells giving. We are grateful to Hongyun Liu and Wei Lan from Institute of Dairy Science, College of Animal Sciences for fluorescence microscope guidance and Dr. Chao Sun from Agricultural, Biological and Environmental Test Center for assistance with flow cytometry, Zhejiang University.

No conflict of interest exists.