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The Journal of Immunology, 2006, 177: 4103-4112.
Copyright © 2006 by The American Association of Immunologists, Inc.

Selective Ablation of Matrix Metalloproteinase-2 Exacerbates Experimental Colitis: Contrasting Role of Gelatinases in the Pathogenesis of Colitis1

Pallavi Garg*, Mauricio Rojas{dagger}, Anupama Ravi*, Katrina Bockbrader*, Steven Epstein*, Matam Vijay-Kumar{ddagger}, Andrew T. Gewirtz{ddagger}, Didier Merlin* and Shanthi V. Sitaraman2,*,{ddagger}

* Division of Digestive Diseases, Emory University, Atlanta, GA 30322; {dagger} Division of Pulmonary, Allergy, and Critical Care Medicine, Department of Medicine, Emory University, Atlanta, GA 30322; and {ddagger} Department of Pathology, Emory University, Atlanta, GA 30322


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The matrix metalloproteinases (MMPs), MMP-2 and MMP-9, share structural and substrate similarities and are up-regulated during human as well as animal models of inflammatory bowel disease. We recently demonstrated that epithelial-derived MMP-9 is an important mediator of inflammation and tissue damage in colitis. In this study, we examined the role of MMP-2 in acute colitis. Colitis was induced using two models, administration of dextran sodium sulfate (DSS) and Salmonella enterica subsp. serovar Typhimurium (S.T.). Bone marrow chimeras were performed using bone marrow cells from wild-type (WT) and MMP-2–/– mice. Colitis was evaluated by clinical symptoms, myeloperoxidase assay, and histology. MMP-2 protein expression and activity were up-regulated in WT mice treated with DSS or S.T. MMP-2–/– mice were highly susceptible to the development of colitis induced by DSS (or S.T.) compared with WT. During inflammation, MMP-2 expression was increased in epithelial cells as well as in the infiltrating immune cells. Bone marrow chimera demonstrated that mucosa-derived MMP-2 was required for its protective effects toward colitis. Furthermore, we demonstrate that severe colitis in MMP-2–/– is not due to a compensatory increase in MMP-9. Finally, we show that MMP-2 regulates epithelial barrier function. In contrast to MMP-9, mucosa-derived MMP-2 may be a critical host factor that is involved in the prevention or cessation of the host response to luminal pathogens or toxins, an important aspect of healing and tissue resolution. Together, our data suggest that a critical balance between the two gelatinases determines the outcome of inflammatory response during acute colitis.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Inflammatory bowel diseases (IBD),3 which include ulcerative colitis and Crohn’s disease, are chronic, incurable intestinal disorders. Ulcerative colitis and Crohn’s disease are characterized by continuous or discontinuous mucosal inflammation, respectively, with inflammatory cell infiltration, epithelial cell destruction, connective tissue defects, and ulceration of the mucosa of the major portion of intestine and/or colon. The etiopathogenesis of IBD is poorly understood, but considered to be multifactorial, involving genetic susceptibility and environmental factors that lead to the synthesis and release of inflammatory cytokines and recruitment of inflammatory cells culminating in tissue destruction. Accumulating data from several studies indicate the involvement of matrix metalloproteinases (MMPs) in the pathogenesis of IBD via their influence on the function and migration of inflammatory cells, mucosal ulceration, as well as matrix deposition and degradation (1, 2, 3). Clinical and experimental studies have demonstrated an increase in the abundance of these matrix-degrading proteases in IBD, and inhibition of MMP activation has been shown to improve experimental colitis (4, 5, 6, 7).

MMPs are a family of Zn2+-dependent extracellular matrix-degrading endopeptidases that share common functional domains and activation mechanisms (8, 9, 10). MMPs are the only mammalian enzymes capable of catalyzing cleavage of the interstitial collagen, types I, II, III, or IV, in their native triple helical form, at the neutral pH of the extracellular space (11). Depending on substrate specificity, amino acid similarity, and identifiable sequence modules, the MMPs can be classified into four major subgroups: collagenases (MMP-1, -3, -8), gelatinases (MMP-2, -9), stromelysins (MMP-3, -7, -10, -12), and membrane-type metalloproteinases 1 through 5. Although MMPs play important roles in normal tissue remodeling, dysregulated expression has been implicated in several pathological processes, such as arthritis (1, 12), atherosclerosis, myocardial infarction (13), colorectal cancer, tumor invasion (14, 15), and IBD (4, 16). In many acute and chronic inflammatory events such as IBD, MMP levels have been shown to be up-regulated (17, 18, 19, 20, 21, 22, 23).

Among the MMPs, MMP-2 and MMP-9 are the two known gelatinases that degrade denatured collagen and are consistently up-regulated during active flares of IBD in human as well as in animal models of colitis (17, 18, 23, 24, 25, 26). Gelatinases differ from other MMPs in terms of their structure as well as substrate specificity. Both MMP-2 and MMP-9 have the prototypic structure that is present in other MMPs: signal peptide, followed by the prodomain, catalytic domain, hinge region, and the hemopexin domain, respectively (1). However, they differ from other MMPs by the presence of fibronectin-like repeats in their catalytic domain. Both MMP-2 and MMP-9 share common substrates and are expressed by similar cell types (1, 11). A major structural difference between MMP-2 and MMP-9 is the lack of an additional type V collagen-like domain in MMP-2 (27). Little is known regarding the role of the MMP-2 or MMP-9 in inflammation or injury of colon during IBD.

We recently demonstrated that MMP-9 activity and protein expression were absent from normal colonic mucosa, but were up-regulated during experimental colitis in response to luminal toxin (dextran sodium sulfate; DSS) as well as bacteria Salmonella enterica subsp. serovar Typhimurium (S.T.) (24). MMP-9–/– mice exposed to DSS or S.T. had significantly reduced inflammation and mucosal injury and were protected against acute colitis. Immune response to systemic administration of S.T. was also not affected in MMP-9–/– mice. Finally, epithelial- but not immune cell-derived MMP-9 was required for tissue damage. The role and function of MMP-2 in the pathogenesis of intestinal inflammation are not known. In this study, we sought to characterize the role of MMP-2 in the pathogenesis of experimental acute colitis.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Experimental animals

The Animal Care Committee of Emory University approved all procedures performed on animals and the procedures were in accordance with the Guide for the Care and Use of Laboratory Animals, published by the U.S. Public Health Service. MMP-2–/– mice (C57BL/6 background) were a gift from L. Matrisian (Vanderbilt University, Nashville, TN). The homozygous MMP-2-deficient mice used were progeny of heterozygous breeding pairs of C57BL/6 background with disruption of the MMP-2 gene that were backcrossed for more than six generations (28). These mice developed normally and were fertile. To confirm the absence of MMP-2 gene expression, genomic DNA was isolated and the disruption of the MMP-2 gene was confirmed via PCR using primers designed to specifically detect hetero- and homozygote mice. For MMP-2 wild type (WT), we used primers 5'-CAACGATGGAGGCACGAGTG-3' and 5'-CCGGGGAACTTGATCATGG-3' (29). For MMP-2–/– mice, we used primer 5'-TGCAAAGCGCATGCTCCAGA-3' and primer 5'-TGTATGTGATCTGGTTCTTG-3' (29). As described by Perez et al. (29), PCR conditions used were: one cycle of 94°C for 5 min, followed by 34 cycles of 94°C for 1 min, 59°C for 1 min, 72°C for 1 min and 30 s, and one cycle of 72°C for 7 min. The lack of MMP-2 protein and activity was confirmed by Western blot and gelatin zymography, respectively, among MMP-2–/– mice. Age- and sex-matched WT and MMP-2–/– littermates used in the study were between 6 and 8 wk old at the beginning of the experimental protocol and were maintained under conditions as described previously (24).

Induction of DSS colitis

Colitis was induced in two groups of age- and sex-matched male and female WT and MMP-2–/– littermates, by oral administration of DSS (ICN Biomedicals) at 3% (w/v) in tap water ad libitum for 6 days. Age-matched male and female WT and MMP-2–/– littermates receiving tap water served as control. Mice were observed daily and evaluated for changes in body weight and development of clinical symptoms.

S.T. infection

Gut-restricted S.T. infection was induced, as described previously (24, 30). To prepare S.T. inocula, bacteria (S.T. SL3201) were grown overnight at 37°C in 10 ml of Luria-Burtani broth in a 20-ml container with shaking (150 rpm) and were then used to inoculate fresh medium (1:100) and were grown under the same conditions for 2–3 h until an OD at 550 nm of 0.35–0.6 was reached. Bacterial cultures were then diluted in normal saline, and the CFU were enumerated by plating a dilution series of the inoculum. Water and food were withdrawn 4 h before treating with 7.5 mg of streptomycin (75 µl of sterile water containing streptomycin or 75 µl of sterile water by gavage). Afterward, animals were supplied with food and water ad libitum. At 20 h after streptomycin treatment, food and water were withdrawn again for 4 h before mice were infected with 108 CFU of S.T. (50-µl suspension in PBS) or treated with vehicle. Thereafter, food and water were offered immediately. Mice were sacrificed by CO2 inhalation, and tissue samples were processed, as described for the DSS colitis model (6).

Gelatin zymography

The activity of MMPs is measured by zymography under nonreducing conditions, as described previously (24, 31). Briefly, snap-frozen samples of colon were homogenized with homogenizer and extracted in ice-cold nonreducing extraction buffer (20 µl/mg tissue). After centrifugation, the supernatant was collected and the total protein concentration was measured by Lowry method using protein assay reagent (Bio-Rad). Protein from tissue samples was electrophoretically separated on 7.5% polyacrylamide agarose gel copolymerized with gelatin (1.5 mg/ml) as a substrate. The gels were washed three times for 15 min each with 2.5% Triton X-100 (Sigma-Aldrich) to remove the SDS and to allow the electrophoresed enzymes to renature, before being incubated in zymography buffer (5 mmol/L CaC12 and 50 mmol/L Tris-HC1 (pH 7.5)) for 18 h at 37°C. The gels were then stained with 0.5% Coomassie brilliant blue R-250 (Bio-Rad) and destained with methanol:acetic acid:water (v/v 4:1:5). Prestained standard high-range protein markers (Bio-Rad) were used to determine the molecular weights of the gelatinases. Clear digested regions representing MMP activity were quantified using an imaging densitometer (Camera Imaging Densitometer; Model 8300; Alpha Innotech).

Protein extraction and Western blot analysis

As described previously (24), for Western blot analysis, colon tissues obtained as above were homogenized and extracted with lysis buffer. Samples were then centrifuged at 12,000 rpm for 10 min at 4°C and the resulting supernatant was used for assays. The total protein concentration of all samples was measured by Lowry method using protein assay reagent (Bio-Rad). A total of 40 µg was boiled for 5 min in Laemmli’s sample buffer (Bio-Rad) and electrophoresed in 10% SDS-PAGE gels. Proteins were transferred to nitrocellulose (Bio-Rad), and the membrane was then blocked in 5% nonfat dry milk for 1 h. Incubation was performed overnight at 4°C with Abs for tissue inhibitor of metalloproteinase-1 (TIMP-1) (1:1000), TIMP-2 (1:1000), MMP-2 (5 µg/ml), and MMP-9 (1:2000) (Chemicon International). Subsequently, the membranes were washed with Tris-NaCl-Tween 20 and incubated with a goat anti-rabbit (1:2500) or goat anti-mouse (1:4000) IgG HRP conjugate (Bio-Rad) for 1 h at room temperature. Membranes were developed with Western Lightning Chemiluminescence Reagent Plus (PerkinElmer) and quantified by image analysis (32).

Clinical activity score

Assessment of body weights, stool consistency, and the presence of occult/gross blood by a guaiac test (Hemoccult Sensa; Beckman Coulter) were determined daily for each mouse. Colitis was quantified with a clinical score, as described by Cooper et al. (33), using the parameters of weight loss, stool consistency, and fecal blood. Briefly, no weight loss was considered as 0 points, weight loss of 1–5% was scored 1 point, loss of 5–10% as 2 points, 10–20% weight loss as 3 points, and a loss of >20% of the weight was scored as 4. The stool character was characterized as normal (0), soft with well-formed pellets (1), soft without pellets (2), or diarrhea (4). For occult blood, no blood was scored 0, positive hemoccult scored as 2 points, and gross bleeding was scored 4. The total score was added to get a clinical activity score ranging from 0 to 12. Six days after the induction of colitis, mice were euthanized by CO2/hypothermia. The abdominal cavity was exposed by a midline laparotomy, and the entire colon was removed from the caecum to the anus. The colon was flushed with cold PBS and opened longitudinally for morphologic studies. The length and weight of the colon were measured, and tissue obtained from each colon was processed for further assays.

Histological assessment of colitis

Colonic specimens obtained as above were fixed in formalin and coded for blind microscopic assessment of mucosal lesions (descending colon for DSS colitis and caecum for S.T. colitis). Sections were stained with H&E. Microscopic sections were analyzed and histologic scoring was performed, as described by Cooper et al. (33), based on three variables, according to the severity of the induced damage. Briefly, for inflammation, rare inflammatory cells in the lamina propria were counted as 0; increased numbers of granulocytes in the lamina propria as 1; confluence of inflammatory cells extending into the submucosa as 2; and a score of 3 was given for transmural extension of the infiltrate. For crypt damage, intact crypt was scored 0, loss of 1/3 basal counted as 1, loss of 2/3 basal was counted as 2, entire crypt loss was scored as 3, change of epithelial surface with erosion as 4, and a score of 5 was given for confluent erosion. For evaluation of ulcers, an absence of ulcer was scored 0, 1–2 foci of ulcerations were scored as 1, 3–4 foci of ulcerations were scored as 2, and confluent/extensive ulceration was scored 3. These values were added to give a total histological score of 11.

Myeloperoxidase (MPO) activity in the colon

Neutrophil infiltration into colon was quantified by measuring MPO activity, as described previously (24, 34). Briefly, a portion of colon was homogenized in 1:20 (w/v) of 50 mM phosphate buffer (pH 6.0) containing 0.5% hexadecyltrimethyl ammonium bromide (Sigma-Aldrich), on ice using a Polytron homogenizer. The homogenate was sonicated for 10 s, freeze-thawed three times, and centrifuged at 14,000 rpm for 15 min. Supernatant (14 µl) was added to 1 mg/ml o-dianisidine hydrochloride (Sigma-Aldrich) and 0.0005% hydrogen peroxide, and the change in absorbance at 460 nm was measured. One unit of MPO activity was defined as the amount that degraded 1 µmol peroxidase per minute at 25°C. The results were expressed as absorbance per gram of tissue.

Immunofluorescence

Colon tissue from WT and MMP-2–/– mice embedded in paraffin was obtained, as described by Castaneda et al. (24). The sections were rehydrated using graded alcohols. Sections were treated with 0.5% Triton X-100 + 0.08% saponin in PBS at room temperature for 35 min. The sections were rinsed in PBS and incubated with rhodamine-phalloidin (1/60) diluted in PBS for 40 min at room temperature. Sections were subsequently blocked with 2% BSA for 1 h at room temperature. Sections were incubated with primary Ab, anti-MMP-2 (Chemicon International; 15 µg/ml in 2% BSA in PBS solution), or rabbit IgG (Sigma-Aldrich; control), or anti-actin (Sigma-Aldrich; 1:50,000 in 2% BSA) for 1 h at room temperature. After washing three times with PBS, they were incubated with FITC-conjugated anti-rabbit secondary Ab (Bio-Rad; 1/100 in 2% BSA in PBS solution) or rhodamine-conjugated anti-mouse secondary Ab (Bio-Rad; 1/100 in 2% BSA in PBS solution), for 45 min at room temperature and then mounted with Slow Fade (Molecular Probes) mounting medium and examined using Zeiss LSM microscope.

Bone marrow transplantation

Bone marrow transplantation was performed, as described previously (24, 35). Briefly, the femur and tibia were removed and stripped off all muscle and sinew, and bone marrow cells were harvested from both femurs and tibias by flushing the bone cavity with basal marrow medium (Iscove’s medium; Cambrex). After washing with PBS, bone marrow cells were resuspended in basal marrow medium. Approximately 5 x 106 cells in 50 µl were transplanted retro-orbitally. Four treatment groups with 10 animals per group were used (WT->WT, WT->MMP-2–/–, MMP-2–/–->WT, and MMP-2–/–->MMP-2–/–). Mice were given neomycin at 2 mg/ml for the first week of posttransplantation, after which they were switched to tap water. Engraftment was verified by genotyping bone marrow cells using specific primers (29), as described previously in this work. Five weeks after transplantation, we induced colitis by 3% DSS and mice were assessed daily for rectal bleeding, weight loss, and diarrhea, and the clinical score was obtained. At the end of the experimental period (6 days for DSS) (24), mice were sacrificed and colons were processed for histology, MPO assay, and immunohistochemistry. Engraftment was verified by performing genotyping on the bone marrow cells at the end of 4 wk after transplantation.

In vivo permeability

In vivo permeability assay to assess barrier function was performed using an FITC-labeled dextran method, as described (36). Briefly, 8- to 10-wk-old WT and MMP-2–/– mice were used. Food and water were withdrawn for 4 h and mice were gavaged with permeability tracer (60 mg/100 g body weight of FITC-labeled dextran, m.w. 4000; Sigma-Aldrich). Serum was collected retro-orbitally; fluorescence intensity of each sample was measured (excitation, 492 nm; emission, 525 nm; Cytofluor 2300; Millipore, Waters Chromatography); and FITC-dextran concentrations were determined from standard curves generated by serial dilution of FITC-dextran. Permeability was calculated by linear regression of sample fluorescence (Excel 5.0; Microsoft).

Statistical analysis

The data are presented as mean ± SEM. Statistical analysis was performed using GraphPad Instat 3 software (<www.graphpad.com>). Groups were compared using nonparametric tests, and significance of the differences between groups was assessed using the Mann-Whitney U test or the Wilcoxon signed-ranks test for paired data. Values of p < 0.05 were considered statistically significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
MMP-2 expression and activity are induced during DSS colitis in mice

DSS colitis model, a rapid and reproducible model of colitis that mimics human IBD in several respects, was used to investigate the expression of MMP-2 during colitis. WT C57BL/6 mice were administered 3% DSS in their drinking water (w/v) for 6 days. Mice were weighed daily, and stool was examined for consistency and the presence of blood. Mice were euthanized after 6 days, and protein lysates were prepared from the colon for zymography and Western blot analysis, as described in Materials and Methods. Zymography was performed using a gelatin-impregnated PAGE to evaluate gelatinolytic activity. A representative zymogram and its densitometric analysis are shown in Fig. 1A. Pro-MMP-2 activity was observed in colonic extracts prepared from WT mice that had been fed with water (Fig. 1A, lanes 1–4). There was no active MMP-2 detected in these mice. In contrast to mice fed water, colonic extracts from WT animals given DSS showed strong induction of pro- and active MMP-2 activity (pro, 2 ± 0.3-fold and active, 7.5 ± 3.9-fold over water-treated mice, respectively; mean ± SE, 6 mice/group; Fig. 1A, lanes 5–9). MMP-2 activity steadily increased over the course of DSS administration starting on day 1 (data not shown).


Figure 1
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  		FIGURE 1. MMP-2 protein activity and expression are up-regulated during DSS colitis. C57BL/6 mice were given water or 3% DSS (w/v) for 6 days, at which time mice were euthanized. Colon was harvested and processed for gelatin zymography (A) and Western blot (B). A, Representative zymogram of protein (30 µg/lane) from the colon of an individual WT mouse with (A, lanes 5–9) or without (A, lanes 1–4) DSS treatment shows gelatinolytic band in each lane. Zymogram was quantitated by scanning densitometry, and densities relative to vehicle-treated controls are graphed. Values are representative of three individual experiments, mean ± SE; n = 5 for the DSS-treated groups, and n = 4 for the control group in each of the experiments. B, Representative Western blot of protein from the colon of WT given water (lanes 1–4) or DSS (lanes 5–8) probed with anti-MMP-2 (5 µg/ml) (Chemicon International). Each lane shows protein (30 µg/lane) from an individual mouse with or without DSS treatment. Western blots were quantitated by scanning densitometry, and densities relative to water-treated controls are graphed. Values are representative of three individual experiments, mean ± SE; n = 5 for the DSS-treated groups, and n = 4 for the control group in each of the experiments. Significantly different from mice given water; *, p < 0.05; **, p < 0.001.

 
Western blot analysis of colonic protein extracts from WT mice treated with water or DSS was performed to confirm the identity of gelatinolytic activity observed on zymography. Animals that had not been exposed to DSS showed constitutive MMP-2 protein expression (Fig. 1B, lanes 1–4). Marked induction of MMP-2 protein expression was observed in colitic samples (Fig. 1B, lanes 5–8). WT mice exhibited immunoreactivity at 72 kDa consistent with pro-MMP-2, while the colitic samples showed immunoreactivity at a slightly lower molecular mass at 62 kDa consistent with active MMP-2. Densitometric analysis revealed 8.2 ± 4.1-fold increase in MMP-2 protein expression in extracts from mice exposed to DSS compared with mice fed water (mean ± SE, 6 mice/group; p < 0.01).

The proteolytic activity of MMPs is regulated by the balance between zymogen activation and enzyme inhibition through their endogenous inhibitors, {alpha}-macroglobulins and TIMPs (16). Given the importance of TIMPs on the biological activities of MMPs, we next examined whether the expression of TIMP-1 and TIMP-2 was influenced differentially by DSS administration. Fig. 2 shows the expression levels of TIMP-1 and TIMP-2 in WT mice fed with DSS or water. TIMP-1 levels were unchanged (Fig. 2), while TIMP-2 levels were decreased in mice treated with DSS (Fig. 2, lanes 5–8). Together, these data suggest that MMP-2 activity and protein expression are induced during DSS colitis along with a significant decrease in TIMP-2.


Figure 2
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  		FIGURE 2. TIMP-2, but not TIMP-1, expression is down-regulated during DSS colitis. C57BL/6 mice were given water or 3% DSS (w/v) for 6 days, at which time mice were euthanized. Colon was harvested and processed for Western blot. Representative Western blot of protein from the colon of WT given water (lanes 1–4) or DSS (lanes 5–8) probed with anti-TIMP-1 and anti-TIMP-2 (1:1000) (Chemicon International), respectively. Each lane shows protein (30 µg/lane) from an individual mouse with or without DSS treatment. Western blots were quantitated by scanning densitometry, and densities relative to water-treated controls are graphed. Values are representative of three individual experiments, mean ± SE; n = 5 for the DSS-treated groups, and n = 4 for the control group in each of the experiments. Significantly different from mice given water; *, p < 0.05.

 
MMP-2–/– mice developed severe inflammation in response to DSS and were unable to recover from DSS-induced colitis

To further investigate the role of MMP-2 in the pathogenesis of colitis, we used C57BL/6 mice with targeted deletion of MMP-2 (28). These mice exhibit normal phenotype. We administered DSS in drinking water to age (8–10 wk)- and sex-matched C57BL/6 WT and homozygous MMP-2–/– mice. The mice were compared for the clinical signs of disease, including weight changes, stool consistency, and occult blood, according to grading system previously described (24, 33). Both WT and MMP-2–/– mice exposed to DSS developed signs of colitis within 6 days after the administration of DSS. However, colitis was significantly worse in MMP-2–/– mice given DSS, as evidenced by a clinical disease activity score of 7.9 ± 1 and 11.1 ± 0.5, respectively (mean ± SE, 6 mice/group; p < 0.01; Fig. 3A). Almost all of the MMP-2–/– mice developed diarrhea after day 3. These mice were hemo-occult positive starting from day 2 and exhibited frank bleeding on day 5 after DSS administration. DSS-induced colitis is characterized by the presence of inflammation of the colon manifested by crypt destruction, mucosal damage, epithelial erosions, and infiltration of inflammatory cells into the mucosal tissue. Tissues collected from WT and MMP-2–/– mice exposed to DSS were examined histologically and compared with those from normal controls. As shown in Fig. 3, B and C, a mean histological score of 9.7 ± 0.7 was observed in MMP-2–/– mice given DSS compared with a mean score of 6.8 ± 0.6 in WT mice given DSS (mean ± SE, 6 mice/group; p < 0.01 compared with MMP-2–/– mice). Interestingly, ulceration was significantly increased in MMP-2–/– mice. The ulcers were not only more in number, but involved larger surface area compared with WT mice (Fig. 3C). Histological signs of inflammation were not detected in the water control groups (data not shown). Thus, the data obtained corroborated the results obtained from clinical analysis and confirmed the deleterious effect of targeted MMP-2 deletion toward the development of colitis. These data also suggest a potential role of MMP-2 as a protective MMP against colitis.


Figure 3
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  		FIGURE 3. MMP-2–/– develop severe DSS-induced colitis. Eight- to 10-wk-old WT C57BL/6 and MMP-2–/– mice were weighed and given water or DSS (3% w/v) for 6 days. Mice were euthanized, and colon was processed for histology. Disease severity was assessed and expressed in terms of clinical activity score in A. Clinical score was calculated using parameters of stool consistency, weight loss, and fecal blood. Each bar represents mean ± SE; *, p < 0.02. B, mean histological score; *, p < 0.02 and #, p < 0.05 compared with WT. C, representative sections from two individual experiments. n = 5 for DSS group, and n = 4 for control group. Colon specimens from descending colon were fixed in formalin, and histological score was calculated on H&E sections based on three variables: inflammatory cells, crypt damage, and ulcers. Both WT and MMP-2–/– mice developed inflammatory infiltration and crypt damage. However, MMP-2–/– mice showed extensive ulceration compared with WT mice.

 
To monitor the effect of lack of MMP-2 on the healing phase of colitis, we observed mice for 1 more additional wk after DSS treatment, during which time animals were given water. All of the MMP-2–/– mice died during this period, while only 25% of the WT mice died. Fig. 4A shows the percentile of change in body weight during healing phase and indicates that change in body weight of MMP-2–/– was 2-fold more than WTs. Fig. 4B shows the survival curve of WT and MMP-2–/– mice, indicating higher mortality among WTs compared with knockouts during recovery phase. These data implicate the importance of MMP-2 in intestinal recovery and healing following acute colitis.


Figure 4
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  		FIGURE 4. MMP-2–/– mice exhibit impaired recovery from DSS-induced colitis. WT and MMP-2–/– mice were given water or DSS for 6 days, as described in Fig. 3 (n = 4 per group). After 6 days, mice were given water and followed for weight loss (A) and mortality (B) during recovery phase. Each bar represents mean ± SE; #, p < 0.05.

 
MMP-2 mice exhibit increased susceptibility to Salmonella-induced colitis

As an alternate model of colitis, we used oral infection with S.T., during which S.T. is administered after pretreatment of mice with streptomycin. In this model, S.T. induces clinical and histological features of enterocolitis predominantly involving the caecum (24, 30). We chose this model because it recapitulates some aspects of clinical and histological human infection as well as acute flares of IBD, wherein mucosal-pathogen interaction is thought to play an important role in the pathogenesis. The characteristic histological feature of gut-restricted S.T. enteritis includes neutrophil infiltration of the intestinal mucosa, the hallmark of infectious colitis, as well as acute flares of IBD. Other histological features of S.T. colitis include epithelial ulceration, edema, and induction of ICAM-1. WT and MMP-2–/– mice were pretreated with streptomycin and then administered S.T. Mice were sacrificed at 24 and 48 h after the administration of S.T., and colonic tissue was removed, weighed with the contents, and photographed. The caecum was processed for histology and MPO activity. The caeca of all the mice infected with S.T. appeared pale and shriveled to a small size and were filled with purulent exudates. MMP-2 activity was significantly up-regulated in WT mice treated with S.T. (Fig. 5A, lanes 3 and 4) similar to that seen in WT mice during DSS colitis. MMP-2 activity was not detected in the MMP-2–/– mice with (Fig. 5A, lanes 7 and 8) or without (Fig. 5A, lanes 5 and 6) treatment of S.T. MPO activity reflected the clinical observation (Fig. 5B). MMP-2–/– mice showed significantly increased MPO activity compared with WT mice. Clinical symptoms and histological changes appeared at 24 h in WT and MMP-2–/– mice. However, MMP-2–/– mice showed increased severity of inflammation compared with WT mice at 48 h (Fig. 5C) as marked by leukocyte infiltration with loss of crypts as well as ulcerations at 48 h in MMP-2–/– mice treated with S.T. Taken together, these data suggest that MMP-2 is up-regulated during S.T.-induced colitis and MMP-2–/– mice develop severe S.T.-induced colitis compared with WT mice.


Figure 5
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  		FIGURE 5. MMP-2–/– mice showed increased susceptibility to Salmonella-induced colitis. WT and MMP-2–/– mice (C57BL/6) were pretreated with streptomycin before the administration of S.T. and sacrificed 48 h after the administration of S.T. The caecum and colon were harvested. Western blot analysis (A) of protein from the caecum of WT given water (lanes 1 and 2) or S.T. (lanes 3 and 4) and MMP-2–/– given water (lanes 5 and 6) or S.T. (lanes 7 and 8) probed with anti-MMP-2 was done. Each lane shows protein (40 µg/lane) from an individual mouse with or without S.T. treatment. beta-actin served as the loading control. Colons were snap frozen in liquid nitrogen, and MPO activity was measured (B) as an index of neutrophil infiltration into the injured tissue. WT mice are represented by {blacksquare}, and MMP-2–/– mice by Figure 5. Each bar represents mean ± SE. n = 3 animals for each group. *, p < 0.005. C, H&E sections based on three variables: inflammatory cells, crypt damage, and ulceration showed that both WT and MMP-2–/– mice developed inflammatory infiltration and crypt damage to S.T. induced colitis though knockouts showed more ulceration compared to WT.

 
MMP-2 localizes to epithelial cells and immune cells

We next determined the localization of MMP-2 in normal and inflamed colon. Immunofluorescence of colon sections performed with affinity-purified MMP-2 Ab directed against the catalytic domain of mouse MMP-2 confirmed the presence of this enzyme mainly in epithelial cells (Fig. 6A). There was no staining in the lamina propria immune cells (Fig. 6A, boxed). In mice fed with DSS, MMP-2 staining was strongly induced and was seen in the crypt epithelial cells (Fig. 6B). Strong MMP-2 staining was also detected in the infiltrating immune cells (Fig. 6C). Control section stained with isotype Ab (anti-rabbit IgG) instead of primary Ab showed no staining. There was no staining visualized in MMP-2–/– mice (Fig. 6D). Thus, the expression of MMP-2 is induced in mice exposed to DSS and is localized to epithelial and immune cells.


Figure 6
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  		FIGURE 6. MMP-2 localizes to the epithelial and immune cells. C57BL/6 mice were weighed and randomized into four groups: WT water or DSS (3% w/v) and MMP-2–/– water or DSS (3% w/v) for 6 days. Mice were euthanized, and colon was processed for immunofluorescence using MMP-2-specific Ab (Chemicon International; 15 µg/ml). Immunoreactive MMP-2 was revealed by FITC-conjugated secondary Ab. Shown here are representative sections from three individual experiments. n = 5 for DSS group, and n = 4 for control group. MMP-2 staining in WT mice fed water (A), WT mice fed 3% DSS showing epithelial staining (B), and infiltrating immune cells (C). Boxed area in A, lamina propria immune cells. MMP-2–/– mice showed no staining (D).

 
Mucosa-derived MMP-2 mediates its protective effect

To understand the role of MMP-2 in the pathogenesis of colitis, we first determined the relative contribution of mucosal vs immune cell-derived MMP-2 in the development of colitis. To explore this question, we generated bone marrow chimeras of WT or MMP-2 mice. Four treatment groups with 10 animals per group (D:WT->R:WT, D:WT->R:MMP-2–/–, D:MMP-2–/–->R:WT, D:MMP-2–/–->R:MMP-2–/–; D, donor; R, recipient) were included. Bone marrow transplantation was done, as described in Materials and Methods. All mice recovered uneventfully from bone marrow transplantation, except 20% of the MMP-2–/–->MMP-2–/– group died. Five weeks after bone marrow transplantation, mice were fed 3% DSS in drinking water. The mice were compared for the clinical signs of weight loss, stool consistency, and occult blood, and mice were sacrificed after 6 days of DSS. Colitis was worst in all MMP-2–/– mice whether they received WT or MMP-2–/– bone marrow (Fig. 7, A and B). These mice had extensive crypt damage, epithelial ulceration, crypt abscess formation, and infiltration of inflammatory cells and neutrophils in the mucosa (Fig. 7C). Fig. 7A shows the clinical score of bone marrow chimeras, indicating that MMP-2–/– mice receiving bone marrow cells from MMP-2–/– or WT mice had higher scores (9.1 ± 0.3 and 11.4 ± 0.4, respectively) compared with WT mice receiving bone marrow cells from WT or MMP-2–/– mice (6.4 ± 0.5 and 7.1 ± 0.5, respectively). MPO assay (Fig. 7B) reflected the same fact that MMP-2–/– mice that received MMP-2–/– or WT bone marrow exhibited significantly increased MPO activity and obvious signs of colonic inflammation and tissue damage compared with WT mice that received WT or MMP-2–/– bone marrow. To verify the efficiency of bone marrow reconstitution, we performed immunofluorescence staining of DSS-treated chimeric mice. As shown in Fig. 7D, MMP-2–/– recipients of WT bone marrow showed MMP-2 staining restricted to immune cells, while WT recipient of MMP-2–/– bone marrow showed staining restricted to the epithelium. These data suggest that reconstitution of the transplanted bone marrow was efficient. Collectively, these data demonstrate that immune cell-derived MMP-2 is neither required for migration nor sufficient to induce tissue damage in DSS-induced colitis. Furthermore, these data suggest that mucosal MMP-2 expression during colitis is required for its protective effect on the development of colitis.


Figure 7
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  		FIGURE 7. Bone marrow chimeras showed that mucosa-derived MMP-2 mediates its protective effect. Bone marrow transplantation was performed, as described in Materials and Methods (D, donor; R, recipient; and KO, MMP-2–/– mice). Mice were euthanized, and colon was processed for assessment of colitis, MPO assay, histology, and immunofluorescence. A, Clinical score showing that KO mice had the worst colitis whether they have received WT or KO bone marrow. Each bar represents mean ± SE. n = 10, a or c vs b: p < 0.01; c vs a: p < 0.05. B, Shows the measure of MPO activity that is an index of neutrophil infiltration into the injured tissue, and was performed from colons of bone marrow chimeras, as described in Materials and Methods. C, Histological assessment of colitis after administration of 3% DSS for 5 days, indicating extensive crypt damage, epithelial ulceration, crypt abscess formation, and infiltration of inflammatory cells and neutrophils in the mucosa of KO mice receiving bone marrow from WT or KO mice compared with WT mice receiving bone marrow cells from WT or KO mice. Top panel, x20 magnification; middle panel, x40 magnification; and lower panel, x60 magnification. D, Immunofluorescence staining of DSS-treated chimeric mice using MMP-2-specific Ab. Immunofluorescent MMP-2 was revealed by rhodamine-conjugated secondary Ab. Sections were counterstained with anti-actin and revealed by FITC-conjugated secondary Ab.

 
Severe colitis in MMP-2–/– mice is not due to a compensatory increase in MMP-9

We recently demonstrated that MMP-9 is up-regulated during experimental colitis and mucosa-derived MMP-2 mediates tissue damage during colitis. MMP-9–/– are protected from DSS- and S.T.-induced colitis (24). To verify whether a compensatory increase in MMP-9 among MMP-2–/– mice is contributing to the severity of colitis seen in these mice, we performed Western blot in the colonic tissue lysates of WT and MMP-2–/– mice administered water or DSS. Fig. 8, A and B, shows a representative Western blot of WT and MMP-2–/–, respectively. In contrast to WT mice fed water (Fig. 8A, lanes 2–4), colonic extracts from WT mice given DSS (Fig. 8A, lanes 5 and 6) showed increased MMP-9 expression. Interestingly, MMP-9 expression was not increased in MMP-2–/– mice fed with water (Fig. 8B, lanes 2–4) and administration of DSS increased the expression of MMP-9 (Fig. 8B, lanes 5 and 6) similar to WT mice given DSS. These data suggest that a compensatory increase in MMP-9 does not play a role in the severity of colitis seen with the deletion of MMP-2.


Figure 8
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  		FIGURE 8. Severe colitis in MMP-2–/– mice is not due to a compensatory increase in MMP-9 expression. WT (A) and MMP-2–/– (B) mice (C57BL/6) were given water or 3% DSS (w/v) for 6 days, at which time mice were euthanized. Colon was harvested and processed for Western blot. Representative Western blot of protein from the colon of WT and MMP-2–/– mice given water (lanes 2–4) or DSS (lanes 5 and 6) probed with anti-MMP-9 is shown in A and B, respectively. Purified MMP-9 (4 ng/lane) was loaded in lane 1 in both A and B. Each lane shows protein (40 µg/lane) from an individual mouse with or without DSS treatment. beta-actin served as the loading control.

 
MMP-2–/– mice exhibit compromised epithelial barrier function

Because our data show that epithelial-derived MMP-2 plays a protective role in the development of colitis, we hypothesized that MMP-2 may play a role in epithelial barrier function and barrier function may be decreased in MMP-2–/– mice, rendering them susceptible to injury and inflammation. Loss of barrier function provided by epithelial cells is thought to be the initial inciting event that underlies injury and inflammation in many intestinal disorders, including IBD (37, 38). Such barrier defects result in the migration of antigenic material, previously confined to the intestinal lumen, into the submucosa, exposing lamina propria immune cells to naive Ags, eliciting inflammatory response and epithelial injury that characterize these diseases (37, 38). To test our hypothesis, we studied barrier function in WT and MMP-2–/– mice using a FITC-labeled dextran method, as described in Materials and Methods. Mice were administered FITC-dextran by gavage, and fluorescence was quantitated in the serum at 4 and 24 h after the administration of FITC-dextran. As shown in Fig. 9, WT mice showed an FITC-dextran of 1.05 ± 0.1 mg of FITC/µg protein/h. In comparison, there was ~2-fold increase in FITC-dextran levels in MMP-2–/– mice (1.8 ± 0.03 mg of FITC/µg protein/h), suggesting decreased barrier function in these mice.


Figure 9
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  		FIGURE 9. MMP-2–/– mice exhibit compromised epithelial barrier function. WT and MMP-2–/– mice (C57BL/6) were gavaged FITC-dextran (m.w. 4000), as described in Materials and Methods. Serum was obtained retro-orbitally and processed for fluorescence. Data are represented as mg FITC/µg protein/hour. Each bar represents mean ± SE. n = 10; p < 0.01.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
In this study, we demonstrate that although MMP-2 and MMP-9 are both gelatinases that originate from similar epithelial sources as well as share structural and substrate similarities, they have opposing effects on the development of acute colitis. Consistent with other studies, we show that MMP-2, like MMP-9, is up-regulated during colitis (4, 16, 24, 39). Using two models of acute colitis, chemical- and bacteria-induced colitis, we demonstrate that MMP-2–/– mice are highly susceptible to the development of colitis compared with their WT counterparts. These results are in contrast to the effect of MMP-9 on colitis; MMP-9–/– are protected from the development of chemical- or bacteria-induced colitis. These results were surprising given that MMP-2 and MMP-9 belong to the family of gelatinases that share structural and matrix substrate similarities. Both MMP-2 and MMP-9 possess the fibronectin repeat characteristic of the gelatinases. However, MMP-2 lacks the collagen V-like domain present in MMP-9, potentially changing their nonmatrix substrate specificity. For example, McQuibban et al. (40) reported contrasting effect of MMP-2 and MMP-9 on MCP-3. MMP-2 degrades MCP-3, while MMP-9 has no effect on the protein. The authors extrapolated the difference in MCP-3 degradation between MMP-2 and MMP-9 to the collagen V domain. The effect of MMP-2 on MCP-3 degradation and possibly degradation of other chemoattractants is thought to underlie the protective effect of MMP-2 on inflammatory arthritis, myocardial damage, and corneal injury (11, 41, 42, 43, 44, 45, 46, 47, 48, 49). In addition to degradation of chemoattractants or chemokines, a role for MMP-2 in tissue remodeling has been shown in wounded corneal epithelial cells. In this study, MMP-9 impaired remodeling, while MMP-2 was required for proper tissue remodeling and wound healing. These authors further showed that fibroblast-derived MMP-2 was required for stromal remodeling that is integral to corneal wound healing. Along with the observations that myofibroblasts are an important source of MMP-2 during intestinal inflammation (19) and are integral to epithelial wound healing, it is likely that MMP-2 may play a role in tissue remodeling that is required for proper wound healing (50, 51).

Our data show that cellular constituents of intestinal mucosa, including epithelial cells as well as immune cells, express MMP-2 in response to DSS or S.T. These observations are consistent with published data that MMP-2 is highly expressed in the intestinal epithelia during human IBD. However, to our knowledge, the role of MMP-2 in the pathogenesis of colitis has not been demonstrated. We demonstrate that MMP-2 may be involved in the regulation of epithelial barrier function. MMP-2–/– mice exhibited barrier dysfunction, as evidenced by increased FITC-dextran translocation in these mice compared with their WT counterpart. Data from human and animal studies have demonstrated a central role for epithelial dysfunction in the pathogenesis of intestinal inflammation (37, 38). The current paradigm for the pathogenesis of IBD supported by evidence from human and animal studies as well as cultured cell models proposes three key components to be necessary for the initiation and progression of disease: 1) disruption of the epithelial barrier (direct effect on barrier or impaired healing in response to injury); 2) access of luminal contents to the lamina propria, that is, immune cells; and 3) an abnormal immune response (37, 38). Epithelial dysfunction is seen in patients much earlier than histologic or clinical manifestation of IBD. It is thought that barrier dysfunction allows contents of the intestinal lumen to mix freely with the contents of the lamina propria, eliciting an immune/inflammatory response, which characterizes inflammatory diseases of the intestine. Our data show that barrier integrity is compromised in MMP-2–/– mice, which may possibly lead to their increased susceptibility to luminal toxins (DSS) or pathogens (S.T.).

There are several possible mechanisms by which MMP-2 might affect barrier function. One possibility is that MMP-2 can module tight junction directly by associating with tight junction proteins. A series of studies have shown that MMP-2 intimately associates with claudins (52, 53, 54). For example, it has been demonstrated that MMP-2 localizes to tight junctions in Madin-Darby canine kidney cells and ectodomain of claudin-1 interacts with the catalytic domain MMP-2 (52). Claudin family proteins are the major constituents of tight junction strands, which are directly involved in paracellular sealing (barrier function) as well as in membrane domain differentiation (fence function) in epithelial and endothelial cell (55, 56). Hence, the association of MMP-2 with claudin may modulate paracellular permeability. In support of this association, our data show that overexpression of MMP-2 increases barrier function measured using FITC translocation (our unpublished data). In this context, it is interesting that MMP-9, unlike MMP-2, has been associated with increased paracellular permeability by cleaving extracellular domain of occludin while having no effect on claudin (57). Such disparate effect of MMP-2 and MMP-9 on permeability may account for their opposite effects on the pathogenesis of colitis.

Given our previous observation that MMP-9 mediates inflammatory response and tissue damage, a possible explanation for the severe colitis seen in MMP-2–/– mice could be related to a compensatory increase in MMP-9 in MMP-2–/– mice. Such a compensatory increase in MMP-9 has been shown to mediate severe inflammatory response in an autoimmune encephalitis model in MMP-2–/– mice (45). The potential role of MMP-2 as a modulator of MMP-9 expression via the membrane-type 1-MMP/TIMP-2 complex has been suggested (45, 58). However, we did not observe any increase in MMP-9 expression or activity in MMP-2–/– mice fed water, and the increase in MMP-9 was modest after DSS colitis compared with WT mice fed DSS. Thus, expression of MMP-9 may be partially modulating the development of colitis in MMP-2–/– mice, while the loss of MMP-2 expression may play a crucial role in the severity of colitis. However, further studies are required to delineate the precise mechanism by which MMP-2 modulates its protective effects.

In summary, we demonstrate that MMP-2, a gelatinase that shares structural and substrate similarities with MMP-9, plays a protective role toward the development of acute colitis. We also show that mucosal MMP-2 contributes to its protective effect, and immune cell-derived MMP-2 is not required for its migration or protective effect. Lastly, we show that MMP-2 is required for intact epithelial barrier function. Together, our data suggest that MMP-2 plays a protective role in the development of murine colitis possibly by contributing to barrier function. Furthermore, a critical balance between the two gelatinases determines the outcome of inflammatory response during acute colitis. Our data are relevant in designing gelatinase-based therapeutic strategies for the treatment of intestinal inflammation.


   Acknowledgments
 
We thank Dr. Lynn Matrisian (Vanderbilt University, Nashville, TN) for providing us C57BL/6 MMP-2–/– mice, and Dr. J. Xu (Emory University, Division of Pulmonary, Atlanta, GA) for helping with bone marrow chimeras.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK06411, Crohn’s and Colitis Foundation of America Senior Research Award and Elseivier Research Initiative Award (to S.V.S.), National Institute of Diabetes and Digestive and Kidney Diseases Grant DK061941 and DK071594 (to D.M.), and Digestive Disease Research Center Grant 5R24DK064399-02. Back

2 Address correspondence and reprint requests to Dr. Shanthi V. Sitaraman, Division of Digestive Diseases, Room 201-F, 615 Michael Street, Whitehead Research Building, Emory University, Atlanta, GA 30322. E-mail address: ssitar2{at}emory.edu Back

3 Abbreviations used in this paper: IBD, inflammatory bowel disease; DSS, dextran sodium sulfate; MMP, matrix metalloproteinase; MPO, myeloperoxidase; S.T., Salmonella enterica subsp. serovar Typhimurium; TIMP, tissue inhibitor of metalloproteinase; WT, wild type. Back

Received for publication April 14, 2006. Accepted for publication June 19, 2006.


   References
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 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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