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Intestinal epithelial cell (IEC) apoptosis contributes to the development of ulcerative colitis (UC), an inflammatory bowel disease (IBD) that affects the colon and rectum. Therapies that target the inflammatory cytokine TNF have been found to inhibit IEC apoptosis in patients with IBD, although the mechanism of IEC apoptosis remains unclear. We therefore investigated the role of p53-upregulated modulator of apoptosis (PUMA), a p53 target and proapoptotic BH3-only protein, in colitis and IEC apoptosis, using patient samples and mouse models of UC. In UC patient samples, PUMA expression was elevated in colitis tissues relative to that in uninvolved tissues, and the degree of elevation of PUMA expression correlated with the severity of colitis and the degree of apoptosis induction. In mice, PUMA was markedly induced in colonic epithelial cells following induction of colitis by either dextran sulfate sodium salt (DSS) or 2,4,6-trinitrobenzene sulfonic acid (TNBS). The induction of PUMA was p53-independent but required NF-κB. Absence of PUMA, but neither absence of p53 nor that of another BH3-only protein (Bid), relieved DSS- and TNBS-induced colitis and inhibited IEC apoptosis. Furthermore, treating mice with infliximab (Remicade), a clinically used TNF-specific antibody, suppressed DSS- and TNBS-induced PUMA expression and colitis. These results indicate that PUMA induction contributes to the pathogenesis of colitis by promoting IEC apoptosis and suggest that PUMA inhibition may be an effective strategy to promote mucosal healing in patients with UC.
The inflammatory bowel diseases (IBDs) ulcerative colitis (UC) and Crohn disease (CD) are thought to result from aberrant activation of the intestinal mucosal immune system (1). Although the pathogenesis of IBD remains unclear, a number of studies have suggested a role of abnormal apoptosis in the intestinal epithelial cells (IECs), resulting from increased cytokine production, such as TNF, IL, and interferon family members (2). Increased IEC apoptosis has been detected at the acute inflammatory sites in UC (3, 4) and CD patients (5). Induction of IEC apoptosis has also been described in a number of studies using murine colitis models (6–8). IEC apoptosis can disrupt intestinal mucosal integrity and barrier function and lead to other changes associated with colitis (9, 10). Furthermore, anti-TNF therapies for treating IBD patients were found to inhibit IEC apoptosis (11, 12). However, the molecular basis of IEC apoptosis in response to intestinal inflammation remains poorly understood.
p53-upregulated modulator of apoptosis (PUMA), a BH3-only Bcl-2 family member, was identified as a downstream target of the tumor suppressor p53 and a potent inducer of apoptosis in diverse tissues and cell types (13). The expression of PUMA is elevated in response to different stimuli through p53-dependent or -independent transcription (14). PUMA binds to antiapoptotic Bcl-2 family members to activate the proapoptotic members Bax and Bak to trigger mitochondrial dysfunction. This results in the release of several apoptogenic mitochondrial proteins, such as cytochrome c and SMAC, leading to caspase activation and cell death (15, 16). Knockout of PUMA in human colon cancer cells or in mice leads to resistance to apoptosis induced by a number of stimuli, such as DNA damage, kinase inhibition, oncogene activation, and endoplasmic reticulum stress (13).
An emerging role of PUMA in gastrointestinal inflammation and tissue injury has been revealed by several recent studies. PUMA is directly activated by NF-κB, a key player in intestinal inflammation (17), in response to TNF-α treatment (18). TNF-α–induced apoptosis in several tissues, including small intestine and colon, is significantly reduced in PUMA-deficient (PUMA-KO) mice (18). Lack of PUMA significantly compromised IEC apoptosis and intestinal damage induced by γ-irradiation (19). Using an acute ischemia/reperfusion model, we previously demonstrated that PUMA is necessary for IEC apoptosis and tissue injury induced by intestinal ischemia/reperfusion, which induces an inflammatory response (20). Furthermore, PUMA was found to be an important regulator of apoptosis in the immune system, and its expression is subject to regulation by immune modulators, such as glucocorticoids and growth hormones, and by bacterial or viral infection (13).
We hypothesized that PUMA is involved in the pathogenesis of colitis by mediating IEC apoptosis induced by intestinal inflammation. We found that PUMA is induced in patients with UC and mouse experimental colitis models, and its induction is correlated with severity of colitis. Our findings suggest that PUMA induction by TNF through NF-κB contributes to colitis and IEC apoptosis, whereas its suppression by anti-TNFs attenuates colitis and IEC apoptosis.
To study the role of PUMA in UC and its associated IEC apoptosis, we first analyzed 23 colitis specimens and 17 uninvolved colonic tissues from patients with UC, including 12 matched pairs from the patients without pancolitis. Signs of colonic inflammation, including submucosa edema and lymphocyte infiltration, were evident in the colitis tissues (Supplemental Figure 1A; supplemental material available online with this article; doi: 10.1172/JCI42917DS1). TUNEL staining revealed apoptosis induction in every colitis specimen compared with uninvolved samples (Figure (Figure1A1A and Supplemental Figure 1, A–C). Apoptosis induction was confirmed by staining for active caspase-3, a biochemical marker of apoptosis (Figure (Figure1B1B and Supplemental Figure 1, A, D, and E). Costaining with the epithelial marker cytokeratin and TUNEL or active caspase-3 detected apoptosis in both epithelial and nonepithelial cells (Figure (Figure1C1C and Supplemental Figure 1D).
PUMA protein expression was found to be elevated in 39.1% of colitis samples (9 out of 23 samples) compared with that in uninvolved tissues by Western blot analysis (Figure (Figure1D1D and Supplemental Figure 2A). PUMA mRNA levels were also found to be induced in colitis specimens relative to those in uninvolved tissues by RNA in situ hybridization (ISH; Figure Figure1E1E and Supplemental Figure 2B) and by real-time RT-PCR analysis (Qiu et al., unpublished observations). Immunostaining for PUMA confirmed its induction in IECs in which apoptosis could be detected simultaneously (Supplemental Figure 2, C and D). Remarkably, elevated PUMA expression significantly correlated with the severity of colitis, with 7.9-fold median PUMA induction in samples with moderate or severe colitis, compared with 3.3-fold induction in those with mild colitis (Figure (Figure1F).1F). Furthermore, induction of apoptosis was also found to correlate with the severity of colitis (Figure (Figure1F)1F) and with PUMA induction (Supplemental Figure 2E).
The above observations prompted us to use mouse models to further investigate the role of PUMA in colitis. Upon treating WT mice with 5% dextran sulfate sodium salt (DSS), an inducer of colitis and colonic inflammation (21), we found a number of inflammatory cytokines to be induced in the colonic mucosa, including Il1a, Il1b, Il6, Tnfa, Icam1, and Vcam1 (Figure (Figure2A).2A). Similarly, PUMA mRNA levels were increased by 8 fold within 24 hours after DSS treatment (Figure (Figure2,2, B and C). The induction of PUMA mRNA by DSS was confirmed by ISH analysis (Figure (Figure2D).2D). PUMA protein was also substantially induced after treatment (Figure (Figure2E2E and Supplemental Figure 3A) and was found to be colocalized with cytokeratin (Supplemental Figure 3B). Furthermore, increased PUMA expression could be detected in purified IECs from DSS-treated mice (Supplemental Figure 4, A and B). In addition to PUMA, BH3-only proteins Noxa and Bim, along with antiapoptotic proteins Bcl-2 and Bcl-XL, were also induced by DSS (Supplemental Figure 4C). Thus, PUMA induction in IECs appears to constitute a rapid response in DSS-induced colitis and colonic inflammation.
DSS-induced colitis is characterized by body weight loss, presence of occult or gross blood per rectum, and tissue damage (21). The disease phenotypes can be measured by the disease activity index, which is the sum of body weight, diarrhea, and rectal bleeding scores (22). To determine whether the induction of PUMA contributes to colitis, WT and PUMA-KO mice were subjected to DSS treatment. Although all DSS-treated mice developed colitis starting from day 3 (Supplemental Figure 5, A and B), the disease activity index score was significantly lower in PUMA-KO mice than in WT mice (Figure (Figure3A).3A). Fewer and smaller colonic ulcers were detected in PUMA-KO mice compared with WT mice after DSS treatment for 7 days (Figure (Figure3,3, B–D). Histological analysis revealed substantially less epithelial damage and disruption of crypt architecture in PUMA-KO mice than in WT mice (Figure (Figure3,3, E and F). However, similar levels of inflammatory cell infiltration were found in both WT and PUMA-KO mice (Figure (Figure3E).3E). Induction of myeloperoxidase (MPO) activity, which is an indicator of neutrophil infiltration, was also similar in WT and PUMA-KO mice after DSS treatment (Supplemental Figure 5C). These results suggest that PUMA contributes to DSS-induced colitis and intestinal damage after the initiation of the inflammatory response.
To determine whether PUMA is involved in DSS-induced colitis through induction of apoptosis, we compared apoptosis induction in DSS-treated WT and PUMA-KO mice. DSS treatment induced DNA laddering, a classical marker of apoptosis, in the mucosa of WT mice but not in that of PUMA-KO mice (Figure (Figure4A).4A). DNA fragmentation quantified by diphenylamine assay was decreased from 22.9% in WT mice to 8.8% in PUMA-KO mice (Supplemental Figure 6A). Apoptosis detected by TUNEL staining started as early as 1 day after DSS treatment (Supplemental Figure 6B), primarily in the crypt areas (Qiu et al., unpublished observations). Apoptosis in these areas was sustained throughout the course of colitis and was blocked by 60%–80% in PUMA-KO mice (Figure (Figure4,4, B and C, and Supplemental Figure 6B). Cytokeratin and TUNEL double staining confirmed the epithelial origin of the PUMA-dependent apoptosis in the crypt areas (Figure (Figure4D4D and Supplemental Figure 6C). DSS treatment for 7 days also induced apoptosis in the submucosa areas, including regions containing infiltrating lymphocytes, but this apoptosis seemed to be PUMA independent (Figure (Figure4B). 4B).
PUMA promotes apoptosis by activating caspases through a mitochondrial pathway (13). DSS treatment of WT mice led to cytosolic accumulation of cytochrome c, a mitochondrial apoptogenic protein (Figure (Figure4E).4E). The release of cytochrome c was decreased in PUMA-KO mice (Figure (Figure4E).4E). Concordantly, activation of caspase-3, a key caspase in the execution of apoptosis, was significantly reduced in PUMA-KO mice, as shown by the abundance of the caspase-3 cleavage fragment (Figure (Figure4F)4F) and by the analysis of caspase-3 activity (Supplemental Figure 6D). Collectively, these results indicate that PUMA mediates IEC apoptosis in DSS-induced colitis through the mitochondrial pathway.
To verify the role of PUMA in colitis and IEC apoptosis, we analyzed an independent colitis model induced by 2,4,6-trinitrobenzene sulfonic acid (TNBS), which resembles CD (23). Similar to the DSS treatment, TNBS treatment also induced PUMA mRNA and protein in IECs of WT mice (Figure (Figure5,5, A–C, Supplemental Figure 3, A and B, and Supplemental Figure 4, A and B). TNBS treatment induced similar levels of MPO activity and expression of Il1a, Il6, and Tnfa in both WT and PUMA-KO mice (Supplemental Figure 7, A and B). TNBS-induced colitis and colonic damage were substantially decreased in PUMA-KO mice compared with those in WT mice (Figure (Figure5,5, D–F). TNBS-induced IEC apoptosis was also markedly reduced in PUMA-KO mice (Figure (Figure5,5, G and H, and Supplemental Figure 6E). These results confirm the critical role of PUMA in colitis and its associated IEC apoptosis.
PUMA is a downstream target of p53 and plays a critical role in p53-mediated apoptosis (13). WT and p53-deficient (p53-KO) mice were therefore analyzed to determine the role of p53 in DSS-induced and PUMA-mediated colitis and IEC apoptosis. After DSS treatment, the expression of p53 was unchanged in WT mice, and the induction of PUMA remained intact in p53-KO mice (Figure (Figure6A).6A). Deficiency in p53 did not protect mice from DSS-induced body weight loss, diarrhea, and rectal bleeding (Figure (Figure6B).6B). Tissue damage measured by the histological score, and the number of ulcers was also unchanged in p53-KO mice (Figure (Figure6C6C and Supplemental Figure 8, A–C). Similar levels of IEC apoptosis and caspase-3 activation were detected in DSS-treated WT and p53-KO mice (Figure (Figure6D6D and Supplemental Figure 8, D–F). Furthermore, TNBS-induced PUMA expression, colitis, and IEC apoptosis were also found to be completely p53 independent (Figure (Figure6,6, E and F, and Supplemental Figure 9, A–C). Therefore, the effects of PUMA on colitis and IEC apoptosis were not attributable to p53. To further determine the potential role of other BH3-only proteins in colitis, mice deficient in the BH3-only protein Bid (Bid-KO mice) were analyzed. We found that colonic damage and IEC apoptosis induced by DSS or TNBS were not different in WT and Bid-KO mice (Supplemental Figure 10, A–H).
Our recent study showed that PUMA can be induced by TNF-α through an NF-κB–dependent but p53-independent mechanism (18). This prompted us to test whether NF-κB and TNF are involved in PUMA induction in DSS- and TNBS-induced colitis. Indeed, administration of the NF-κB inhibitor, Bay 117082, blunted the induction of PUMA in DSS-treated WT mice, in conjunction with reduced activation (phosphorylation) of the p65 subunit of NF-κB (Figure (Figure7A).7A). Similarly, pretreatment with infliximab (Remicade), an anti-TNF antibody clinically used for IBD treatment, also suppressed DSS-induced PUMA expression and p65 phosphorylation (Figure (Figure7B7B and Supplemental Figure 11A), suggesting that the induction of PUMA by DSS is mediated by TNF through NF-κB. In addition to PUMA induction, TNF inhibition, an important means for treatment of IBD (1), suppressed DSS-induced colitis (Figure (Figure7C),7C), colonic damage (Figure (Figure7D7D and Supplemental Figure 11B), ulcer formation (Supplemental Figure 11C), and IEC apoptosis (Figure (Figure7,7, E and F). The extent of suppression conferred by TNF inhibition was similar to that observed in PUMA-KO mice.
Consistent with the findings from the DSS model, the induction of PUMA by TNBS was also reduced following NF-κB inhibition (Figure (Figure8A).8A). Cotreatment with infliximab compromised TNBS-induced PUMA expression and p65 phosphorylation (Figure (Figure8B),8B), colonic damage (Figure (Figure8C8C and Supplemental Figure 12A), and IEC apoptosis (Figure (Figure8D8D and Supplemental Figure 12, B and C). Furthermore, pretreatment with the TNF inhibitor pentoxifylline also inhibited colitis and IEC apoptosis induced by DSS (Supplemental Figure 13, A–I) and those induced by TNBS (Supplemental Figure 14, A–E). However, NF-κB inhibition by pretreating mice with Bay 117082 did not protect against DSS- or TNBS-induced colitis and IEC apoptosis (Figure (Figure9).9). Collectively, these results suggest that inhibition of PUMA expression contributes to the antiinflammatory effects of TNF inhibitors.
Apoptotic cell death has been implicated as a major homeostatic and pathogenic mechanism of intestinal epithelium (2). In this study, we found that increased PUMA expression is correlated with the severity of colitis and induction of apoptosis in human UC samples. PUMA deficiency in mice abrogated DSS- and TNBS-induced colitis and IEC apoptosis, supporting the hypothesis that PUMA functions as a critical mediator of IEC apoptosis and a significant modulator of UC. The functional role of PUMA in IBDs needs to be validated using other cohorts of patient samples. The broader implications of our study can be verified using other colitis models, such as the IL10-deficient genetic model. Whether PUMA-mediated IEC apoptosis is responsible for disrupting the barrier function of the intestinal epithelial monolayer remains to be determined. PUMA has recently been shown to regulate T cell apoptosis (24). It is possible that PUMA may contribute to colitis by affecting T cell apoptosis, which is known to be abnormally regulated in IBDs (25, 26). In addition to PUMA, other BH3-only proteins, such as Noxa and Bim, may also be involved in colitis (27).
The induction of PUMA mRNA and protein occurred within 12–24 hours after DSS or TNBS treatment and could be detected throughout the treatment. This rapid and sustained PUMA induction seems to be a downstream inflammatory response, as cytokine induction and lymphocyte infiltration could still be detected in PUMA-KO mice. The induction of PUMA by DSS and TNBS is driven, at least in part, by TNF-α, which is the major cytokine that initiates and perpetuates intestinal mucosal inflammation and damage (28). Interestingly, Il6 and Tnfa expression was reduced in PUMA-KO mice compared with that in WT mice after TNBS treatment (Supplemental Figure 7A), suggesting a feedback loop in which PUMA-mediated apoptosis further enhances cytokine production. Our previous studies have already shown that deficiency in the p65 component of NF-κB, a key regulator of intestinal inflammation (17), blunted PUMA induction by TNF-α in vitro and in vivo (18). This observation, along with the finding that NF-κB or TNF inhibition suppressed PUMA induction by DSS or TNBS, suggests that the TNF/NF-κB axis is responsible for PUMA induction and subsequent IEC apoptosis in colitis. Potential involvement of other cytokines is also possible, as suggested by a recent study, showing PUMA induction by interferon-γ through the transcription factor IRF-1 (29). Unlike PUMA or TNF inhibition, NF-κB blockage did not alleviate colitis, probably reflecting the ability of NF-κB to activate both proapoptotic proteins, such as PUMA, and antiapoptotic proteins, such as Bcl-2 and Bcl-XL (Supplemental Figure 4C).
p53 is absolutely necessary for PUMA induction in IECs by DNA-damaging agents such as γ-irradiation (19). It also activates several proapoptotic proteins implicated in IBDs, such as Fas, death receptor 4 (DR4), and DR5 (30). Surprisingly, DSS- and TNBS-induced PUMA expression and colitis are completely p53-independent. p53 is also unnecessary for the induction of PUMA in response to intestinal ischemia/reperfusion (20). However, our study was focused on acute changes at early time points, whereas chronic colitis has been shown to induce DNA damage and trigger a p53 response (31). FoxO3A, a transcription factor that induces PUMA in response to cytokine deprivation (32), is not required for PUMA induction by TNF-α (18). Our results seem to reveal what we believe to be a novel apoptotic pathway through the TNF-α/NF-κB/PUMA axis, which may play a broad functional role in tissue inflammatory response and damage. It will be interesting to test whether this pathway is also involved in mucosal inflammation induced by bacterial or viral infection.
Modulation of PUMA expression may contribute to the effectiveness of current IBD treatment regimens, such as anti-TNF (33). TNF inhibitory agents may block IEC apoptosis by inhibiting PUMA expression. Our recent studies show that suppressing PUMA expression using an antisense approach significantly improved animal survival and protected against intestinal damage induced by high doses of γ-irradiation (19). Growth factors, such as insulin-like growth factor 1 and basic fibroblast growth factor, also alleviate radiation-induced intestinal injury, at least in part, by suppressing PUMA (34). Therefore, PUMA inhibition may offer an effective way to protect against IEC apoptosis and thereby may serve as a novel anti-IBD strategy. In conclusion, our study has demonstrated that PUMA contributes to colitis by promoting IEC apoptosis and may be a useful target for developing new IBD treatment options.
Frozen specimens of 23 colitis and 17 uninvolved tissues, including 12 matched pairs of samples from patients with UC without pancolitis, were obtained from the Digestive Disease Tissue Resource of the University of Pittsburgh. The acquisition of tissue samples was approved by the Institutional Review Board at the University of Pittsburgh. Written informed consent was received from each patient prior to inclusion in the study. Twelve males and eleven females were represented in colitis group, with the group age ranging from 10–70 years old, whereas 8 males and 9 females were represented in uninvolved group, with the group age ranging from 20–70 years old. Among the 23 patients in the UC group, colitis was found in the colons of 20 patients, the rectums of 10 patients, the ceca of 2 patients, the duodenum of 1 patient, and the terminal ileum of 1 patient. Among the 12 patients whose matched pairs of samples were analyzed, 5 were treated with steroids, 2 received immunomodulator treatment, and 1 was subjected to anti-TNF therapy.
All animal experiments were approved by the Institutional Animal Care and Use Committee at the University of Pittsburgh. Mice in the C57BL/6 background with different genotypes, including WT, PUMA-KO (PUMA–/–), Bid-KO (Bid–/–), and p53-KO (p53–/–), were generated by breeding and identified by PCR genotyping, as previously described (19, 20, 35). The mice were housed in microisolator cages and allowed access to water and chow ad libitum.
Six- to ten-week-old littermates were treated with either DSS or TNBS to induce colitis. For DSS treatment, mice were fed with 5% DSS (MP Biomedicals) in drinking water. Mice received DSS treatment for 1, 3, 5, or 7 days and were immediately sacrificed after treatment. For TNBS treatment, mice were presensitized on the skin for 1 week with TNBS (Sigma-Aldrich) at 40 mg/kg body weight in 50% ethanol. After fasting overnight, the presensitized mice received 100 mg/kg TNBS by rectal injection with a 1-ml syringe fitted with a catheter, as previously described (36). Sacrifice of mice was performed 3 days after TNBS treatment. For TNF inhibition, mice were i.p. injected at 1 hour prior to DSS or TNBS treatment with the anti-TNF antibody infliximab (Remicade; Centocor Ortho Biotech) at 10 mg/kg or the TNF inhibitor pentoxifylline (Sigma-Aldrich) at 200 mg/kg. For NF-κB inhibition, mice were injected at 1 hour prior to DSS or TNBS treatment with 8 mg/kg of the NF-κB inhibitor BAY 117082 (EMD Biosciences) once daily. After TNF or NF-κB inhibition, mice were treated with DSS or TNBS and then sacrificed using the same schedule as those treated with DSS or TNBS alone.
Preparation of intestinal mucosa scraping and isolation of DNA, total RNA, and protein extracts from mouse tissues were performed as previously described (20, 37). Real-time RT-PCR was performed on a Mini Opticon Real-time PCR System (Bio-Rad) with SYBR Green (Invitrogen). Primers for RT-PCR are listed in Supplemental Table 1, except for the previously described PUMA and β-actin primers (20). Agarose (2%) gel electrophoresis was used to verify PCR products.
After protein extracts were subjected to NuPage gel (Invitrogen) electrophoresis, Western blotting was performed using antibodies for human PUMA (15); murine PUMA (Abcam); p53, Bax, and p65 (all from Santa Cruz Biotechnology Inc.); active caspase-3, Bcl-2, Mcl-1, and cytochrome c (all from BD Biosciences); phospho-p65 (Ser536) and Bad (both from Cell Signaling Technology); Bak (Millipore); Bcl-XL (Imgenex); Bim (EMD Biosciences); Noxa (Merck); Bcl-w (Enzo Life Sciences); Bid (a gift from Xiao-Ming Yin at Indiana University, Indianapolis, Indiana, USA); and β-actin (Sigma-Aldrich) as previously described (15). For mouse analysis, representative results of 2 to 3 mice analyzed are presented, with each lane on a Western blot representing a single animal. Western blot band intensities were quantified using ImageJ software ( http://rsbweb.nih.gov/ij/).
Tissue sections (5-μm thick) were deparaffinized, rehydrated, and treated with 3% hydrogen peroxide, followed by antigen retrieval in boiling 0.1 M citrate (pH 6.0) buffer for 10 minutes once. The sections were then blocked by 20% rabbit serum for 30 minutes. Immunostaining was performed as previously described for active caspase-3 (34). PUMA staining was performed at 4°C overnight using a mouse anti-PUMA antibody (Sigma-Aldrich), with Alexa Fluor 594 (Invitrogen) for signal detection. Cytokeratin staining was done at 4°C overnight using a mouse anti-cytokeratin antibody (Abcam), with Alexa Fluor 488 (Invitrogen) or Alexa Fluor 594 for signal detection. For double staining, TUNEL staining was performed after PUMA or cytokeratin staining. Cytokeratin staining was performed prior to PUMA or active caspase-3 staining. Cells with positive staining were scored in at least 100 crypt sections and reported as mean ± SD.
Probe template was prepared by RT-PCR amplifying a 398-bp PUMA cDNA fragment with one of the primers containing a T7 promoter sequence at 5′. Digoxigenin-labeled (DIG-labeled) RNA probes were transcribed using the DIG RNA Labeling Kit (SP6/T7) (Roche Diagnostics). Paraffin-embedded tissue was sectioned (5-μm thick), rehydrated, denatured with 0.2 N HCl, treated with Proteinase K (Ambion), postfixed in 4% paraformaldehyde, and acetylated. After a 1-hour prehybridization at 60°C, denatured DIG-labeled probes were hybridized to sections at 60°C for 16 to 24 hours. Hybridization buffer contained 65% formamide, 5X SSC, 2% Roche blocking powder, 1 μg/ml yeast tRNA, 50 μg/ml heparin, 5 mM EDTA, and 0.05% 3-([3-cholamidopropyl] dimethylammonio)-1-propanesulfonate. After a number of washes to decrease salt concentration, slides were incubated with α-DIG–horse radish peroxidase (Abcam), biotinyl tyramide (DAKO), α-biotin–horse radish peroxidase (Abcam), and then additional biotinyl tyramide (DAKO) to amplify the signal. After incubation with α-biotin–alkaline phosphatase (Abcam), the signal was visualized using nitroblue-tetrazolium-chloride/5-bromo-4-chlor-indolyl-phosphate (Roche Diagnostics) solution, and counterstaining was applied with nuclear Fast Red (Vector Laboratories). When staining was complete, slides were washed in water and mounted in Clear Mount (Electron Microscopy Sciences).
Dissected mouse colons were cut open longitudinally and incubated in 50-ml tubes with 20 ml Ca2+- and Mg+-free Hank’s Buffered Salt Solution (CMF-HBSS; Invitrogen) containing 10 mM dithiothreitol and 1.5 mM EDTA at 4°C for 1 hour. After incubation, the tube was vigorously shaken for 2 minutes to dislodge IECs. The cell suspensions were passed through a 100-μm cell strainer (BD Biosciences) and were centrifuged at room temperature at 400 g for 5 minutes. The cell pellet was resuspended in 2 ml CMF-HNBSS and incubated at 4°C for 1 hour with agitation with a pre-made mixture of 100 μl TBS, 5 μl rat anti-mouse epithelial cell adhesion molecule (EpCAM), and 50 μl anti-rat IgG Danabeads (Invitrogen). The beads were isolated by a Magnetic Particle Concentrator (Invitrogen) and washed 3 times with CMF-HBSS. The isolated IECs were stained for EpCAM and also lysed for Western blotting.
The murine colitis phenotypes were measured by the disease activity index score, which is the sum of body weight, diarrhea, and bleeding scores determined daily during the treatment, as previously described (22). All scores were relative to the scores on day 1, which were set as 0. For body weight, no weight loss was scored as 0, loss of 1%–5% weight registered as 1, loss of 6%–10% weight registered as 2, loss of 11%–20% weight registered as 3, and loss of higher than 20% weight registered as 4. For stool consistency, 0 was scored for well-formed stool pellets, 2 was scored for pasty and semiformed stools that did not adhere to the anus, and 4 was scored for liquid stools that adhered to the anus. For bleeding, which was analyzed by the Hemoccult fecal occult blood test (Beckman Coulter), 0 was assigned for no blood, 2 was assigned for positive Hemoccult, and 4 was assigned for gross bleeding. All scores were blindly confirmed by 2 trained individuals.
Entire colons were collected from the treated mice and were fixed flat between wet filter papers in 10% neutral buffered formalin for 48 hours. The tissues were stained with 0.2% methylene blue (Sigma-Aldrich) in formalin solution for 5 minutes, and ulcerous areas were counted within 24 hours under a microscope. The tissues were subsequently processed as previously described (19). Paraffin-embedded sections (5-mm thick) were subjected to H&E staining for histological analysis. Histological scores were determined blindly based on the previously described criteria (38): 0, normal; 1, moderate mucosal inflammation without erosion and ulcer; 2, severe mucosal inflammation with erosion; 3, severe mucosal inflammation with ulcer (<1 mm); 4, severe mucosal inflammation with ulcer (1–3 mm); 5, severe mucosal inflammation with ulcer (>3 mm).
DNA laddering and fragmentation were analyzed as described previously (20). TUNEL staining was performed using the ApopTag Kit (Chemicon International) according to the manufacturer’s instructions. TUNEL-positive crypts were counted in 100 randomly selected crypts. Cytochrome c release was analyzed by Western blot probing of mitochondrial and cytosolic fractions isolated from intestinal mucosal scraping by differential centrifugation (20). Caspase-3 activity was measured using a colorimetric assay kit (EMD Biosciences), according to the manufacturer’s instructions, or by Western blotting for active caspase-3.
Statistical analysis was carried out using GraphPad Prism IV software. Student’s t test was used to generate P values, except for the experiments involving repeated measures, which were analyzed using 2-way ANOVA. Differences were considered significant if the probability of the difference occurring by chance was less than 5 in 100 (P < 0.05).
We thank Xiao-Ming Yin at Indiana University for providing Bid-KO mice, our surgeons and research nurses for helping collect the tissue samples, and Crissy Dudgeon and Brian Leibowitz for critical reading. This work is supported by NIH grants CA106348 and CA121105, American Cancer Society grant RSG-07-156-01-CNE, the Career Investigator Award from the American Lung Association/CHEST Foundation (to L. Zhang), the Flight Attendant Medical Research Institute (FAMRI), the Alliance for Cancer Gene Therapy (ACGT), American Cancer Society grant RGS-10-124-01-CCE, and NIH grants CA129829 and U01DK085570 (to J. Yu). L. Zhang is a scholar of the V Foundation for Cancer Research.
Conflict of interest: The authors have declared that no conflict of interest exists.
Citation for this article: J Clin Invest. 2011;121(5):1722–1732. doi:10.1172/JCI42917.
Bin Wu’s present address is: Department of Gastroenterology, The Third Affiliated Hospital of Sun Yat-Sen University, Guangzhou, China.