See “Drugs and Tight Junctions: Adverse Effects and Opportunities for New Therapeutic Approaches” by Terrin et al on page 444.
Mucositis is a common adverse-effect of cancer chemotherapy for which there is no present treatment. Mucosal barrier injury occurs approximately in 40% of patients after standard doses of chemotherapy and in 100% of patients undergoing high-dose chemotherapy (1). Chemotherapy-induced intestinal injury has been described as a consequence of different processes: apoptosis, hypoproliferation, inflammatory response, altered absorptive capacity, and bacteria proliferation and colonization (2–5).
Methotrexate (MTX), an inhibitor of dihydrofolate reductase and DNA synthesis, is widely used in children in cancer chemotherapy, with documented activity against leukemia, lymphoma, breast cancer, and head and neck cancers (6). MTX, at lower doses, is also used as an anti-inflammatory agent for chronic inflammatory diseases such as inflammatory bowel diseases, with gastrointestinal and extraintestinal adverse events (7). Several studies have shown that MTX at high doses causes structural and functional injury to the gastrointestinal tract. Apoptosis, hypoproliferation, inflammatory response, and altered absorptive capacity have already been described (1,3,8). We also recently reported that proteolysis dysregulation could be involved in the occurrence of intestinal mucositis (9–11). These changes clinically manifested by severe enterocolitis, gastrointestinal ulceration, diarrhea, and anorexia (12). MTX treatment in rodent models produces similar intestinal mucosal injury to that seen in humans (13), and is associated with hypoproliferation of enterocytes and a disruption of intestinal barrier function (11). Gut barrier disruption could be related to dysregulation of proliferation/apoptosis balance but also to other mechanisms. Intestinal barrier is regulated in part by tight junctions (TJs), which are multiprotein structures involved in the regulation of cell polarity, proliferation, and differentiation (14). TJs consist of transmembrane proteins and peripheral membrane proteins in which zonula occludens (ZO)-1, occludin, and claudin-1 are well characterized. In intestinal disorders, that is, inflammatory bowel diseases (15), irritable bowel syndrome (16,17), or intestinal infections (18), alterations of TJ proteins have already been reported. In a recent study, an alteration of ZO-1 expression has been described after MTX treatment (19), but the expression of other TJ-associated proteins, that is, claudins or occludin, has not yet been described. In addition, it remains unknown whether MTX has direct or indirect effect on TJ proteins and what underlying mechanisms are involved. Thus, the aim of the study was to investigate the effect of MTX on TJs in in vivo and in vitro models of chemotherapy-induced mucositis.
Animal care and experimentation complied with both French regulation and European Community regulation (Official Journal of the European Community L 358, December 18, 1986), and M.C. authorized by the French government to use animal models (authorization no. 76-107).
Animals, Housing, and MTX Injections
During 1 week, 200- to 250-g male Sprague-Dawley rats (Elevage Janvier, Le Genest St Isle, France) were acclimatized at 25°C with a 12-hour light-dark cycle before study. Rats were given free access to water and standard diet. Rats were subcutaneously injected during the first 3 days (D0, D1, and D2) with 2.5 mg/kg MTX (Teva Pharma, Courbevoie, France) or NaCl solution (0.9%) as control, as previously described (10,11,20). Experiments were carried out for 9 days and rats were euthanized on D1, D4, or D9.
Euthanasia and Tissue Sampling Collection
Rats were deeply anesthetized with pentothal (400 mg/kg intraperitoneally) and transcardially perfused with 20 mL of phosphate buffered saline (PBS; 140 mmol/L NaCl, 3 mmol/L KCl, 8 mmol/L Na2HPO4, 1.5 mmol/L KH2PO4) followed by 30 mL of 4% paraformaldehyde (PFA) in PBS. The segments of jejunum were collected, postfixed for 12 hours in PBS-4% PFA, dehydrated by soaking in 15% and 30% sucrose solutions, embedded in Tissue-Tek (OCT compound; Fisher Scientific, Torrance, CA), and immediately frozen at −80°C for immunofluorescence analysis. Before perfusion of animals, jejunal segments were taken and rinsed with ice-cold PBS, immediately frozen in liquid nitrogen, and stored at −80°C until analysis. For mRNA, Trizol reagent (Invitrogen, Cergy-Pontoise, France) was previously added to collecting tubes.
Immunohistochemistry and Confocal Laser Scanning Microscopy
The applied primary and secondary antibody species (all from Zymed Laboratories, South San Francisco, CA), dilutions, and sources were mouse anti-claudin-1 (1:100 dilution), rabbit anti-occludin (1:50), rabbit anti-ZO-1 (1:100), tetramethyl rhodamine isothiocyanate (TRITC) goat anti-rabbit immunoglobulin (Ig) G (1:300), and fluorescein isothiocyanate (FITC) goat anti-mouse IgG (1:300). The immunofluorescence analysis was carried out on frozen sections of jejunum. Tissue sections (10-μm thick) were cut using a cryostat (Leica Microsystems, Nussloch, Germany) perpendicular to the longitudinal axis of the tissue. The sections were mounted on glass slides and air dried. Nonspecific binding was blocked with PBS containing 1% bovine serum albumin (BSA) or 10% normal goat serum (NGS) for 1 hour at room temperature and then the sections were incubated at 4°C overnight in the same solution supplemented with primary antibodies. After 3 washes in PBS, immunolabeling was revealed by using the appropriate secondary antibodies for 1 hour at room temperature. All of the antibody dilutions were made using PBS containing BSA or NGS with Triton X-100 (Sigma-Aldrich, St Louis, MO). Negative controls were obtained by substituting the first antibodies by PBS solution.
After immunohistochemistry, microphotographs were acquired with a confocal laser scanning microscopy using an SP2 AOBS inverted microscope (Leica Microsystems) with an argon laser (488 nm). Image acquisition was performed using Leica Confocal software for optimal detection of FITC and TRITC with a 63× oil immersion objective (numerical aperture 1.4).
Western Blot Analysis
The applied primary antibody species, dilutions, and sources were mouse anti-claudin-1 (1:1000 dilution, Zymed Laboratories), rabbit anti-occludin (1:1000, Zymed Laboratories), rabbit anti-ZO-1 (1:1000, Zymed Laboratories), and mouse anti-β-actin (1:5000, Sigma-Aldrich). Tissues and cell monolayers were homogenized in CelLytic buffer (Sigma-Aldrich). Following centrifugation for 15 minutes at 12,000g at 4°C, the supernatant was collected, supplemented with antiproteases and phosphatase inhibitors, and stored at −80°C. Protein levels were determined using the Bradford assay (Protein Assay, Bio-Rad, Marne-la-Coquette, France). For claudin-1 and occludin expression, total proteins (25 μg) were separated on a 4% to 12% gradient polyacrylamide gel (Invitrogen). For ZO-1 expression, 50 μg of proteins were separated on 6% Tris-glycine gel. Proteins were then transferred to a nitrocellulose membrane. The membrane was blocked for 1 hour at room temperature with 5% (wt:vol) nonfat dry milk in Tris-buffered saline (10 mmol/L Tris, pH 8; 150 mmol/L NaCl) plus 0.05% (wt:vol) Tween 20 followed by an overnight incubation at 4°C with primary antibodies. After being washed in Tris-buffered saline-1% Tween 20 buffer, the membranes were incubated with appropriate secondary antibodies (Santa Cruz Biotechnology, Santa Cruz, CA) for 1 hour at room temperature. After 3 additional washes, immunocomplexes were revealed using the enhanced chemiluminescence detection system (GE Healthcare, Orsay, France). Protein bands were scanned (ImageScanner III; GE Healthcare) and quantified using ImageQuant TL software (GE Healthcare).
Quantification of TJ mRNA Expression Using Real-time Polymerase Chain Reaction
Mucosal total RNAs were extracted from jejunal segments as described earlier (21). After reverse transcription of 1.5-μg total RNA into cDNA by using 200 U of SuperScript II Reverse Transcriptase (Invitrogen), quantitative polymerase chain reaction was performed by SYBR Green technology on BioRad CFX96 real-time-polymerase chain reaction system (Bio-Rad) in duplicate for each sample. Glyceraldehyde-3-phosphate dehydrogenase was used as the endogenous reference gene. Specific primers were for claudin-1, 5′-AGGTCTGGCGACATTAGTGG-3′ and 5′-TGGTGTTGGGTAAGAGGTTG-3′; occludin, 5′-CACGTTCGACCAATGC-3′ and 5′-CCCGTTCCATAGGCTC-3′; ZO-1, 5′-GTATCCGATTGTTGTGTTCC-3′ and 5′-TCACTTGTAGCACCATCCGC-3′; and for glyceraldehyde-3-phosphate dehydrogenase, 5′-CATCACTGCCACTCAGAAGA-3′ and 5′-AAGTCACAGGAGACAACCG-3′.
Cell Culture and Treatments
Human epithelial intestinal adenocarcinoma cell line Caco-2 clone TC7 were purchased from the American Type Culture Collection (Manassas, VA). Caco-2 cells were used between passage 20 to 50 and were routinely grown at 37°C in a water-saturated atmosphere with 5% CO2 in Dulbecco Modified Eagle's Medium (Eurobio, Courtaboeuf, France) supplemented with 10% heat-inactivated fetal calf serum (Eurobio), 1% nonessential amino acid, antibiotics (10000 U/mL penicillin, 10 mg/mL streptomycin; Sigma-Aldrich) and 2 mmol/L L-glutamine (Sigma-Aldrich) in 75-cm2 tissue culture flasks. Culture medium was changed every 2 days.
Then, 1 × 106 cells were seeded on porous filter (0.4-μm pore size; Millipore, Molsheim, France) in 6-well plates. Briefly, 1 and 4 mL of culture medium were added to the upper and lower chambers, respectively. The Transwell plates were then incubated at 37°C in a water-saturated atmosphere with 5% CO2. The culture media were changed every day. Confluent cell monolayers were obtained within 12 to 14 days after seeding. The transepithelial electrical resistance (TEER) gradually increased and reached a plateau after day 14, indicating the formation of cell monolayer.
Caco-2 monolayers were treated with 2 doses of MTX (1 and 100 ng/mL) at the basolateral side during a 24-hour time period. Interleukin (IL)-1β was also added in the basolateral chamber in some experiments to have a positive control (1 and 10 ng/mL; Sigma-Aldrich).
To investigate mechanisms involved in Caco-2 barrier dysfunction, Caco-2 cells were preincubated with or without the specific signal transduction inhibitors (Sigma-Aldrich) for 1 to 2 hours before MTX treatment: NF-κB inhibitor, caffeic acid phenethyl ester (CAPE) at 5 μmol/L; JNK-inhibitor II at 10 μmol/L; extracellular signal-regulated kinase (ERK) 1 and 2 inhibitors, UO126 at 10 μmol/L and PD98059 at 24 μmol/L; p38 mitogen-activated protein kinase (MAPK) inhibitor SB203580 at 10 μmol/L; and PI3kinase inhibitor wortmannin at 10 μmol/L were used.
Cell Viability Assay
To assess cell viability by staining metabolically active cells 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT) assay was done as previously described (22). Ten microliters of the MTT labeling reagent (Sigma-Aldrich) was added and cells were incubated for 4 hours at 37°C. Solubilization solution was added (300 μL/well), and data were analyzed by spectrophotometry in a microtiter plate reader (Metertech Σ 960, Taipei, Taiwan) at a wavelength of 550 nm. Data were expressed as percentage of control cells.
Apoptosis was assessed by flow cytometry. Briefly, cells were resuspended in 100 μL Annexin V Apoptosis Detection Kit (Pharmingen, San Diego, CA). After the addition of 4 μL of Annexin-V-FITC and 10 μL of propidium iodure (PI), cells were incubated for 20 minutes at room temperature in the dark. Finally, 400 μL of Annexin V buffer kit was added. Every assay was made in duplicate. The flow cytometry analysis was performed in a FACSCalibur (Becton Dickinson, Franklin Lakes, NJ) using a 488-nm argon laser. For each experimental sample, a total of 10,000 cells were counted. Data were analyzed using the CellQuest program (Becton Dickinson) calculating early and late apoptosis levels.
Measurement of TEER
An epithelial volt-ohm meter (Millipore Corporation, Molsheim, France) was used for measurements of Caco-2 monolayer TEER as previously reported (23,24). TEER was calculated after subtraction of the resistance value of the filters alone.
Determination of Paracellular Permeability
As an independent measure of paracellular permeability of Caco-2 monolayers, we examined the passage of paracellular marker FITC-dextran 4 kDa through Caco-2 monolayer. Briefly, FITC-dextran (10 mg/mL, Sigma-Aldrich) was loaded into the apical Transwell filter compartment during the incubation period. After stimulation with MTX or IL-1β, medium samples were collected from the basolateral Transwell compartment at 6 and 24 hours. The fluorescence level was measured in a 96-well fluorescent plate reader (excitation at 485 nm and emission at 535 nm). Values were converted in concentrations of FITC-dextran (grams per liter) using a standard curve.
Assessment of Cytokine Concentrations in Culture Supernatants
Cell culture supernatants were collected from both apical and basolateral sides after stimulation of Caco-2 monolayers by MTX. Cytokine concentrations were evaluated by specific multiplex kits (Fluorokine MAP, R&D Systems, Abingdon, UK) for IL-1β, IL-6, IL-8, interferon (IFN)-γ, and tumor necrosis factor (TNF)-α according to the manufacturer's instructions as previously described (16). Concentrations were expressed as picograms per milliliter.
Filter-grown Caco-2 monolayers were washed twice in cold PBS and fixed in 4% PFA for 20 minutes. The Caco-2 monolayers were then incubated in blocking solution composed of BSA (1%) and NGS (10%) in PBS containing Triton X-100 for 1 hour. Cells were then incubated with primary antibodies anti-claudin-1, anti-occludin, and anti-ZO-1 (Zymed Laboratories) in blocking solution overnight at 4°C. After being washed with PBS, the filters were incubated in TRITC- or FITC-conjugated secondary antibodies (Zymed Laboratories) for 2 hours at room temperature. After a final wash, 1,4-diazabicyclo[2.2.2]octane was used to mount the filters onto the coverslips and cells were observed on an Axiolmager Z1 microscope (Carl Zeiss, Gottingen, Germany). Cytochemical staining for comparative studies was performed in the same experimental session, and images were obtained using identical imaging parameters.
Results, expressed as mean ± standard error mean, were compared using Prism version 5.0 (GraphPad software, San Diego, CA). Normal distribution was checked using Kolmogorov-Smirnov test and equal variances by Bartlett test. When normality and equal variances were confirmed, results were compared using analysis of variance and Tukey posttests. In other cases, Kruskal-Wallis test followed by Dunn multiple comparisons was used. All of the statistical tests were 2-sided and a P value of <0.05 was considered significant.
Effects of MTX Treatment on In Vivo Model of Chemotherapy-induced Mucositis
In the jejunum of MTX-treated rats, histological damage was increased and villus height was markedly reduced at D4 compared with controls rats. These parameters were restored at D9 (online-only Figure 1, http://links.lww.com/MPG/A89).
TJs Protein Expression
In MTX-treated rats, ZO-1, claudin-1, and occludin expression was not modified at D1 compared with control rats (Fig. 1). In contrast, at D4, claudin-1 and occludin expression was significantly reduced (Fig. 1). We also observed a trend for a decrease of ZO-1 expression at D4 but the difference did not reach significance. Interestingly, claudin-1 and occludin expression was restored at D9. These data were in accordance with our previous study showing a marked increase in intestinal permeability at D4 (11).
mRNA Levels for TJs Proteins
There was no difference in the mRNA expression of occludin and ZO-1 between MTX-treated rats at D1, D4, and D9 and control rats; however, claudin-1 mRNA was significantly increased in MTX-treated rats at D4 as compared with control rats (3.2 ± 0.6 vs 0.9 ± 0.1, P < 0.05).
Immunostaining of TJ Proteins
In control rats, ZO-1 (Fig. 2A), occludin (Fig. 2C), and claudin-1 (Fig. 2E) were present between the epithelial cells lining the TJs. In MTX-treated rats, at D4, ZO-1 (Fig. 2B), occludin (Fig. 2D), and claudin-1 (Fig. 2F) expression was altered at the intercellular junctions of epithelial cells.
These data suggest that both TJ protein expression and localization were affected by MTX treatment in rats. To better understand the underlying mechanisms, we studied the effects of MTX on intestinal epithelial cells in vitro by using Caco-2 cell monolayer.
Effects of MTX Treatment on In Vitro Model of Chemotherapy-induced Mucositis
Caco-2 Paracellular Permeability
As shown in Figure 3A, MTX induced a decrease of TEER, independent of the dose; however, changes in TEER may reflect modifications of paracellular permeability but also transcellular ion exchanges. We, thus, assessed whether MTX affects FITC-dextran (4 kDa) passage across Caco-2 cell monolayer. FITC-dextran flux from apical to basolateral sides was significantly induced by the 2 doses of MTX (1 and 100 ng/mL) at 24 hours (Fig. 3B). These results suggest that MTX significantly increased epithelial paracellular permeability in vitro to a similar extent than IL-1β that is well known to affect gut barrier function (25,26). Because MTX is an inhibitor of folate metabolism, we assessed apoptosis, necrosis, and epithelial cell viability.
Cell Viability and Apoptosis
After 24 hours of treatment, the viability of Caco-2 cells was not affected by IL-1β at both 1 and 10 ng/mL and by MTX at 1 ng/mL; however, Caco-2 cell viability was slightly decreased (approximately 11%) after MTX treatment at 100 ng/mL compared with control cells. Therefore, we examined whether MTX or IL-1β induces apoptosis in our model of cells. There was no statistically significant difference in the percentage of apoptotic cells between MTX- and IL-1β-treated Caco-2 cells compared with control Caco-2 cells (data not shown).
TJ Protein Expression and Localization
MTX at 100 ng/mL decreased ZO-1 (Fig. 4A–B) and occludin expression (Fig. 4A, C). Claudin-1 expression remained unchanged after MTX treatment (Fig. 4A, D). The lowest dose of MTX (1 ng/mL) did not affect TJ protein expression.
We, then, analyzed TJ protein distribution (Fig. 5). In control cells, the TJ proteins ZO-1, occludin, and claudin-1 were localized in a chicken wire pattern. At the lowest dose, MTX did not alter TJ protein distribution (data not shown). In contrast, MTX at 100 ng/mL altered ZO-1 and occludin staining as compared with control cells (Fig. 5), but it did not affect claudin-1 distribution (data not shown). These results suggest that changes in paracellular permeability observed after MTX treatment correlate with alterations of occludin and ZO-1 protein expression and localization.
To better understand the underlying mechanisms of MTX treatment, we assessed the effects of several cell-signaling inhibitors on TEER, and TJs protein expression and localization in Caco-2 cells. Because MTX induced alteration of TJs protein expression only at the highest dose, subsequent experiments have been done only with 100 ng/mL of MTX. The MEK1 and 2 inhibitor U0126 reversed significantly the decrease in TEER observed after MTX treatment (Fig. 6). TEER was also restored after pretreatment of Caco-2 cells with NF-κB inhibitor, CAPE, or JNK inhibitor in MTX-treated cells (Fig. 6). The p38 MAPK inhibitor, SB203580, and the PI3K inhibitor, wortmannin, had no effect on TEER after MTX treatment (Fig. 6).
These results were then confirmed on TJs protein expression and localization. Indeed, pretreatment of Caco-2 cells with U0126, CAPE, or JNK inhibitor prevented the decrease of ZO-1 and occludin expression induced by MTX (data not shown) and the alteration of cellular distribution (Fig. 5).
MTX increased IL-1β and lL-8 production on the apical side at 100 ng/mL (online-only Figure 2, http://links.lww.com/MPG/A89). MTX at 1 ng/mL did not affect IL-1β but increased IL-8 production. A trend for an increase in IL-6 production was observed but difference did not reach significance (online only Figure 2, http://links.lww.com/MPG/A89). In contrast, MTX whatever the used dose had no effect on cytokine production on the basolateral side (data not shown). TNF-α and IFN-γ remained undetectable in all of the conditions.
Treatment of cancer with antimitotic agents often induces adverse effects such as weight loss, anorexia, and oral/intestinal mucositis (2). We (9–11,20,27) and others (3,19) have reported that MTX treatment in rats is associated with diarrhea, intestinal damage including villus atrophy, and altered absorption. These alterations could be related to hypoproliferation, increased apoptosis (11), or inflammatory response (9,11). Previous data reported that intestinal permeability was markedly increased during chemotherapy (1,11); however, the mechanisms underlying gut barrier disruption were poorly documented. The integrity of TJs is essential for the maintenance of intestinal barrier function. We thus report in the present study the modifications of TJ proteins after treatment with MTX, an anticancer agent.
Interestingly, we showed that MTX induced significant modification of TJ protein expression and/or cellular distribution in MTX-treated rats. We provide the first evidence that not only ZO-1 but also occludin and claudin-1 proteins were affected after MTX treatment. Our data are in accordance with a study (19) showing an alteration of ZO-1 localization in MTX-treated rats but no significant modification of ZO-1 expression (19). In contrast, we show that both occludin and claudin-1 expression and localization were altered in MTX-treated rats. The loss of these TJ proteins in jejunal mucosa may therefore reflect an impairment of TJ barrier function at the acute stage of MTX-induced mucositis. Claudin-1 and occludin are transmembrane proteins and ZO-1 has been shown to be colocalized with the cytoplasmic end of occludin (28). Thus, these proteins interact with one another to form a complex protein network involved in the regulation of paracellular permeability.
To better understand the underlying mechanisms of TJ alteration, experiments were carried out in an in vitro model of human intestinal epithelial barrier by using filter-grown Caco-2 cells. This model has been used widely to assess either intestinal absorption and metabolism processes (29) or regulation of intestinal permeability after inflammatory stimuli (25,26,30–32). Interestingly, we observed a decrease in TEER and an increase in paracellular permeability with MTX at 1 and 100 ng/mL. In addition, MTX treatment induced an alteration of expression and cellular distribution of occludin and ZO-1, but no modification of claudin-1 expression. Our study provides the first evidence that MTX per se was able to induce an intestinal barrier disruption by TJ protein alterations. To know whether MTX-induced barrier disruption is a direct or indirect effect of MTX on cells, we measured cytokine production that has been reported to affect TJ proteins (25,26,33–36). MTX treatment did not induce IL-1β and IL-8 production on the basolateral side but only on the apical side; IFN-γ and TNF-α remained undetectable. Further studies are warranted because, to our knowledge, there were no data to evaluate the effects of cytokines added on the apical side on barrier function.
The pathways involved in the effects of MTX on TJ alteration deserve discussion. During chemotherapy, NF-κB activation has been reported both in serum and in the intestinal mucosa and may play a key role in the occurrence of mucositis (37). NF-κB activation results in the transcription of genes encoding MAPK, cyclooxygenase 2, and tyrosine kinase signaling molecules, which ultimately results in tissue injury (38). In contrast, in MTX-treated rats, ERK that regulates cell proliferation was inactivated (39). Thus, to elucidate the signaling pathways involved in MTX effects, we assessed the effects of several inhibitors. We showed that not only NF-κB but also MEK1 and 2 and JNK pathways were involved in MTX-induced barrier disruption. Indeed, specific inhibitors for MEK1 and 2, JNK, or NF-κB restored ZO-1 and occludin expression after MTX. In contrast, p38 MAPK and PI3K inhibitors did not prevent MTX effects. Our study is in accordance with previous studies reporting that MEK1 and 2, JNK, and NF-κB pathways may be involved in TJ regulation in extraintestinal or in intestinal tissues in response to inflammatory stimuli. For instance, in Caco-2 cells, IL-1β-induced TJ alteration was mediated by NF-κB activation (25) and the MEKK-1 pathway (26). After osmotic challenge, occludin and ZO-1 expressions were reduced through the JNK pathway (40). In other cell types, occludin expression was reduced by estradiol through the MEK1 and 2 pathways (41) or by osmotic challenge through the JNK pathway (42). Similarly, fibrinogen decreased ZO-1 and occludin expression through the MEK pathway (43). All of these data suggest that NF-κB, MEK1 and 2, and JNK pathways are involved in the regulation of TJ proteins; however, our study shows for the first time that MTX-induced gut barrier disruption was mediated by NF-κB and the JNK and MEK1 and 2 pathways.
A limitation of our study is that TJ proteins were not similarly affected by MTX in the studied models. Indeed, in the in vivo model, MTX induced a reduction of occludin and claudin-1 expression. In contrast, in Caco-2 cells, ZO-1 and occludin expression was affected. This difference could be explained by contributions from innate immune cells and epithelium. Even if the cultured epithelial model does not mimic exactly a whole animal system, the study in Caco-2 cells provided interesting data on the direct effect of MTX on intestinal epithelial cells and on the signaling pathways. Further studies in MTX-treated rats should evaluate the role of NF-κB and MAPK pathways on the occurrence of intestinal barrier disruption.
In conclusion, in an in vivo rodent model of mucositis, the increase in intestinal permeability induced by MTX is related in part to the alteration of TJ proteins, ZO-1, occludin, and claudin-1. The in vitro data suggest that not only NF-κB but also the MEK1 and 2 and JNK pathways are involved in the gut barrier disruption induced by MTX, and consequently these pathways could be therapeutic targets to limit intestinal damage.
1. Keefe DM, Cummins AG, Dale BM, et al. Effect of high-dose chemotherapy on intestinal permeability in humans. Clin Sci (Lond) 1997; 92:385–389.
2. Keefe DM, Brealey J, Goland GJ, et al. Chemotherapy for cancer causes apoptosis that precedes hypoplasia in crypts of the small intestine in humans. Gut 2000; 47:632–637.
3. Carneiro-Filho BA, Lima IP, Araujo DH, et al. Intestinal barrier function and secretion in methotrexate-induced rat intestinal mucositis. Dig Dis Sci 2004; 49:65–72.
4. Bowen JM, Gibson RJ, Tsykin A, et al. Gene expression analysis of multiple gastrointestinal regions reveals activation of common cell regulatory pathways following cytotoxic chemotherapy. Int J Cancer 2007; 121:1847–1856.
5. Stringer AM, Gibson RJ, Logan RM, et al. Chemotherapy-induced diarrhea is associated with changes in the luminal environment in the DA rat. Exp Biol Med (Maywood) 2007; 232:96–106.
6. Asselin BL, Devidas M, Wang C, et al. Effectiveness of high dose methotrexate in T-cell lymphoblastic leukemia and advanced stage lymphoblastic lymphoma: a randomized study by the Children's Oncology Group (POG 9404). Blood 2011;118:874–83.
7. Hausmann J, Zabel K, Herrmann E, et al. Methotrexate for maintenance of remission in chronic active Crohn's disease: long-term single-center experience and meta-analysis of observational studies. Inflamm Bowel Dis 2010; 16:1195–1202.
8. Sukhotnik I, Shehadeh N, Coran AG, et al. Oral insulin enhances cell proliferation and decreases enterocyte apoptosis during methotrexate-induced mucositis in the rat. J Pediatr Gastroenterol Nutr 2008; 47:115–122.
9. Alamir I, Boukhettala N, Aziz M, et al. Beneficial effects of cathepsin inhibition to prevent chemotherapy-induced intestinal mucositis. Clin Exp Immunol 2010; 162:298–305.
10. Boukhettala N, Leblond J, Claeyssens S, et al. Methotrexate induces intestinal mucositis and alters gut protein metabolism independently of reduced food intake. Am J Physiol Endocrinol Metab 2009; 296:E182–E190.
11. Leblond J, Le Pessot F, Hubert-Buron A, et al. Chemotherapy-induced mucositis is associated with changes in proteolytic pathways. Exp Biol Med (Maywood) 2008; 233:219–228.
12. Pico JL, Avila-Garavito A, Naccache P. Mucositis: its occurrence, consequences, and treatment in the oncology setting. Oncologist 1998; 3:446–451.
13. Taminiau JA, Gall DG, Hamilton JR. Response of the rat small-intestine epithelium to methotrexate. Gut 1980; 21:486–492.
14. Shin K, Fogg VC, Margolis B. Tight junctions and cell polarity. Annu Rev Cell Dev Biol 2006; 22:207–235.
15. Kucharzik T, Walsh SV, Chen J, et al. Neutrophil transmigration in inflammatory bowel disease is associated with differential expression of epithelial intercellular junction proteins. Am J Pathol 2001; 159:2001–2009.
16. Coeffier M, Gloro R, Boukhettala N, et al. Increased proteasome-mediated degradation of occludin in irritable bowel syndrome. Am J Gastroenterol 2010; 105:1181–1188.
17. Bertiaux-Vandaele N, Youmba SB, Belmonte L, et al. The expression and the cellular distribution of the tight junction proteins are altered in irritable bowel syndrome patients with differences according to the disease subtype. Am J Gastroenterol 2011;106:2165–73.
18. Chen ML, Pothoulakis C, LaMont JT. Protein kinase C signaling regulates ZO-1 translocation and increased paracellular flux of T84 colonocytes exposed to Clostridium difficile toxin A. J Biol Chem 2002; 277:4247–4254.
19. Hamada K, Shitara Y, Sekine S, et al. Zonula occludens-1 alterations and enhanced intestinal permeability in methotrexate-treated rats. Cancer Chemother Pharmacol 2010; 66:1031–1036.
20. Boukhettala N, Ibrahim A, Claeyssens S, et al. A diet containing whey protein, glutamine, and TGFbeta modulates gut protein metabolism during chemotherapy-induced mucositis in rats. Dig Dis Sci 2010; 55:2172–2181.
21. Leblond J, Hubert-Buron A, Bole-Feysot C, et al. Regulation of proteolysis by cytokines in the human intestinal epithelial cell line HCT-8: role of IFNgamma. Biochimie 2006; 88:759–765.
22. Mosmann T. Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J Immunol Methods 1983; 65:55–63.
23. Ma TY, Hoa NT, Tran DD, et al. Cytochalasin B modulation of Caco-2 tight junction barrier: role of myosin light chain kinase. Am J Physiol Gastrointest Liver Physiol 2000; 279:G875–G885.
24. Ma TY, Nguyen D, Bui V, et al. Ethanol modulation of intestinal epithelial tight junction barrier. Am J Physiol 1999; 276:G965–G974.
25. Al-Sadi R, Ye D, Dokladny K, et al. Mechanism of IL-1beta-induced increase in intestinal epithelial tight junction permeability. J Immunol 2008; 180:5653–5661.
26. Al-Sadi R, Ye D, Said HM, et al. IL-1beta-induced increase in intestinal epithelial tight junction permeability is mediated by MEKK-1 activation of canonical NF-kappaB pathway. Am J Pathol 2010; 177:2310–2322.
27. Boukhettala N, Ibrahim A, Aziz M, et al. A diet containing whey protein, free glutamine, and transforming growth factor-beta ameliorates nutritional outcome and intestinal mucositis during repeated chemotherapeutic challenges in rats. J Nutr 2010; 140:799–805.
28. Willott E, Balda MS, Heintzelman M, et al. Localization and differential expression of two isoforms of the tight junction protein ZO-1. Am J Physiol 1992; 262:C1119–C1124.
29. Gill RK, Saksena S, Tyagi S, et al. Serotonin inhibits Na+/H+ exchange activity via 5-HT4 receptors and activation of PKC alpha in human intestinal epithelial cells. Gastroenterology 2005; 128:962–974.
30. Chavez AM, Menconi MJ, Hodin RA, et al. Cytokine-induced intestinal epithelial hyperpermeability: role of nitric oxide. Crit Care Med 1999; 27:2246–2251.
31. Boivin MA, Ye D, Kennedy JC, et al. Mechanism of glucocorticoid regulation of the intestinal tight junction barrier. Am J Physiol Gastrointest Liver Physiol 2007; 292:G590–G598.
32. Wang F, Schwarz BT, Graham WV, et al. IFN-gamma-induced TNFR2 expression is required for TNF-dependent intestinal epithelial barrier dysfunction. Gastroenterology 2006; 131:1153–1163.
33. Han X, Fink MP, Delude RL. Proinflammatory cytokines cause NO*-dependent and -independent changes in expression and localization of tight junction proteins in intestinal epithelial cells. Shock 2003; 19:229–237.
34. Ma TY, Iwamoto GK, Hoa NT, et al. TNF-alpha-induced increase in intestinal epithelial tight junction permeability requires NF-kappa B activation. Am J Physiol Gastrointest Liver Physiol 2004; 286:G367–G376.
35. Talavera D, Castillo AM, Dominguez MC, et al. IL8 release, tight junction and cytoskeleton dynamic reorganization conducive to permeability increase are induced by dengue virus infection of microvascular endothelial monolayers. J Gen Virol 2004; 85:1801–1813.
36. Tazuke Y, Drongowski RA, Teitelbaum DH, et al. Interleukin-6 changes tight junction permeability and intracellular phospholipid content in a human enterocyte cell culture model. Pediatr Surg Int 2003; 19:321–325.
37. Logan RM, Gibson RJ, Bowen JM, et al. Characterisation of mucosal changes in the alimentary tract following administration of irinotecan: implications for the pathobiology of mucositis. Cancer Chemother Pharmacol 2008; 62:33–41.
38. Sonis ST. The pathobiology of mucositis. Nat Rev Cancer 2004; 4:277–284.
39. Sukhotnik I, Shteinberg D, Ben Lulu S, et al. Transforming growth factor-alpha stimulates enterocyte proliferation and accelerates intestinal recovery following methotrexate-induced intestinal mucositis in a rat and a cell culture model. Pediatr Surg Int 2008;24:1303–11.
40. Samak G, Suzuki T, Bhargava A, et al. c-Jun NH2-terminal kinase-2 mediates osmotic stress-induced tight junction disruption in the intestinal epithelium. Am J Physiol Gastrointest Liver Physiol 2010; 299:G572–G584.
41. Ning L, Kunnimalaiyaan M, Chen H. Regulation of cell-cell contact molecules and the metastatic phenotype of medullary thyroid carcinoma by the Raf-1/MEK/ERK pathway. Surgery 2008; 144:920–924.
42. Tai LM, Holloway KA, Male DK, et al. Amyloid-beta-induced occludin down-regulation and increased permeability in human brain endothelial cells is mediated by MAPK activation. J Cell Mol Med 2010; 14:1101–1112.
43. Patibandla PK, Tyagi N, Dean WL, et al. Fibrinogen induces alterations of endothelial cell tight junction proteins. J Cell Physiol 2009; 221:195–203.