Article - Cell Press

Cell Host & Microbe

Article Helicobacter pylori Dampens Gut Epithelial Self-Renewal by Inhibiting Apoptosis, a Bacterial Strategy to Enhance Colonization of the Stomach Hitomi Mimuro,1 Toshihiko Suzuki,1,6 Shigenori Nagai,3,4 Gabriele Rieder,5 Masato Suzuki,1 Takeshi Nagai,1 Yukihiro Fujita,1 Kanna Nagamatsu,1 Nozomi Ishijima,1 Shigeo Koyasu,3,4 Rainer Haas,5 and Chihiro Sasakawa1,2,3,* 1Department

of Microbiology and Immunology of Infectious Diseases Control, International Research Center for Infectious Diseases Institute of Medical Science, The University of Tokyo, Tokyo 108-8639, Japan 3Core Research for Evolutional Science and Technology (CREST), Japan Science and Technology Agency, Saitama 332-0012, Japan 4Department of Microbiology and Immunology, Keio University School of Medicine, Tokyo 160-8582, Japan 5Max von Pettenkofer Institute for Hygiene and Medical Microbiology, Ludwig Maximilians University, D-80336 Munich, Germany 6Present address: Division of Bacterial Pathogenesis, Graduate School of Medicine, University of the Ryukyus, Okinawa 903-0125, Japan. *Correspondence: [email protected] DOI 10.1016/j.chom.2007.09.005 2Department

SUMMARY

Colonization of the gastric pits in the stomach by Helicobacter pylori (Hp) is a major risk factor for gastritis, gastric ulcers, and cancer. Normally, rapid self-renewal of gut epithelia, which occurs by a balance of progenitor proliferation and pit cell apoptosis, serves as a host defense mechanism to limit bacterial colonization. To investigate how Hp overcomes this host defense, we use the Mongolian gerbil model of Hp infection. Apoptotic loss of pit cells induced by a proapoptotic agent is suppressed by Hp. The ability of Hp to suppress apoptosis contributed to pit hyperplasia and persistent bacterial colonization of the stomach. Infection with WT Hp but not with a mutant in the virulence effector cagA increased levels of the prosurvival factor phospho-ERK and antiapoptotic protein MCL1 in the gastric pits. Thus, CagA activates host cell survival and antiapoptotic pathways to overcome self-renewal of the gastric epithelium and help sustain Hp infection. INTRODUCTION Chronic Helicobacter pylori (Hp) infection of the stomach is a major risk factor for gastritis and gastroduodenal ulcers, as well as for gastric adenocarcinoma and mucosa-associated lymphoid tissue (MALT) lymphoma (Blaser, 1990; Parsonnet et al., 1991, 1994; Peek and Blaser, 2002). The gastric pits are the primary niche for Hp in colonized hosts (Falk et al., 1993; Mahdavi et al., 2002), and the pits undergo rapid turnover, with pit cells self-renewing within as little as 3 days (Karam and Leblond, 1993). The

short turnover period is sustained by the vigorous proliferation of progenitor cells generating at the isthmus and by rapid apoptosis of the superficial pit cells (Karam and Leblond, 1993; Mills et al., 2002; Oh et al., 2005). Apoptosis is a highly conserved and regulated process for eliminating aged, damaged, or infected cells from tissues (Jiang and Wang, 2004), thus serving as an intrinsic mechanism against bacterial and viral infection. Apoptosis is triggered by two major pathways, an intrinsic pathway and an extrinsic pathway. Intrinsic apoptosis is induced by increased permeability of the mitochondrial outer membrane. This permeability leads to cytochrome c (cyt-c) release, apoptosome formation (cyt-c/Apaf-1), and processing of pro-caspase-9 to the mature form. The extrinsic apoptosis pathway is mediated by ligand/receptor binding, which results in the activation of caspase8 (Budd et al., 2006). Caspase-8 directly activates caspase-3 and Bid, and tBid (as a result of cleavage of Bid by caspase-8) stimulates cyt-c release from the mitochondria. Cyt-c forms apoptosomes and activates caspase-9, which in turn causes maturation of caspase-3. The extrinsic and intrinsic pathways merge at caspase-3 and both ultimately result in cell death. Important mediators of intrinsic apoptosis are members of the B cell lymphoma-2 (Bcl2) family, prosurvival proteins (BCL2, BCL2L1, BCL2L2, MCL1, A1), proapoptotic Bax-like proteins (Bax, Bak, Bok), and proapoptotic BH3-only proteins (Bad, Bim, Bmf, Bik, Hrk, Bid, Puma, Noxa) (Strasser, 2005). Myeloid cell leukemia sequence-1 (MCL1), which is widely expressed, plays an important role in the development (Boucher et al., 2000; Opferman et al., 2005) and survival of myeloma cells (Zhang et al., 2002), although its role in epithelial cells is poorly understood. During infection, Hp delivers CagA by a type IV secretion system (TFSS) into gastric epithelial cells (Odenbreit et al., 2000; Segal et al., 1999), where CagA plays a key role in developing various Hp-associated gastric diseases

250 Cell Host & Microbe 2, 250–263, October 2007 ª2007 Elsevier Inc.

Cell Host & Microbe Pivotal Role of CagA to Accelerate MCL1 Expression

(Blaser et al., 1995; Eck et al., 1997; Kuipers et al., 1995). Recent studies have shown that intracellular CagA is phosphorylated by Src family kinases on tyrosine residues within the five repeated EPIYA sequences (PY region) (Asahi et al., 2000; Selbach et al., 2002), and this PY region of CagA is capable of interacting with many host proteins, including SHP2, GRB2, CSK, c-Met, ZO-1, and CRK (Backert and Meyer, 2006). These interactions of CagA with GRB2, CRK, and SHP2 activate the MEK/ERK pathway and lead to gastric epithelial proliferation, cell-cell dissociation, and increased cell motility (Higashi et al., 2002; Mimuro et al., 2002; Suzuki et al., 2005). However, the impact of CagA’s interaction with each of these host proteins on the development of Hp-associated gastric diseases has not been determined. In addition, in vitro studies have indicated that CagA is capable of deregulating transcriptional activity, such as serum responsive element (SRE)-dependent transcription (Hirata et al., 2002), b-catenin (Franco et al., 2005), and nuclear factor of activated T cells (NFAT) (Yokoyama et al., 2005), but the significance of this regulation is also unclear. Many studies have indicated that, during chronic Hp infection, the imbalance between gastric epithelial proliferation and rapid turnover by apoptosis triggers the development of Hp-associated gastric diseases. In vitro studies have shown that Hp contains several apoptosis-inducing factors, including lipopolysaccharide, g-glutamyl-S-transferase, and VacA (Galmiche et al., 2000; Kawahara et al., 2001; Shibayama et al., 2003). Nevertheless, Hp is capable of readily establishing chronic infection of the gastric epithelium, suggesting that Hp can modulate the rapid turnover of superficial gastric cells. In the present study we have induced apoptosis in the Mongolian gerbil stomach by oral administration of Etoposide (Etp), an intrinsic apoptosis inducer known to cause apoptosis of epithelial cells. Using this pharmacological agent to induce gastric superficial pit epithelium apoptosis, we found that Hp can prevent rapid apoptosis of epithelial cells. We also provide convincing in vivo and in vitro evidence supporting the pivotal role of Hp CagA/MCL1 interplay in modulating rapid turnover of pit epithelial cells via MEK/ERK/SRE activation. Importantly, we confirmed that CagA activity was essential not only to suppress apoptosis but also to promote bacterial colonization of superficial pits in the gerbil stomach. RESULTS Hp CagA Suppresses Apoptosis of the Pit Epithelium In Vivo To assess the distribution of apoptotic cells in stomach superficial epithelial cells, gastric sections from Mongolian gerbils were examined by TUNEL, AAA, GS II, and DNA staining. A few spontaneous apoptotic cells were detected only at pits of the corpus and antrum (Figure 1A, left panels). To induce apoptosis in epithelial cells, we administered Etp, an intrinsic apoptosis inducer, into the stomach of gerbils. Fourteen hours posttreatment, the number of apoptotic cells in the pits increased as indicated by greater exfoliation of mucosal layers (Figure 1A, right

panels). To investigate whether Hp infection affects apoptosis in the gerbil stomach, gerbils not treated with Etp were inoculated with WT, DcagA (CagA-deficient), or DvirB7 (TFSS-deficient) Hp derived from NCTC 11637 or ATCC 43504 strains (Figure S1), and the number of apoptotic epithelial cells in the pits was quantitated 3 and 8 weeks after infection. Although Hp infection slightly increased the number of apoptotic cells in the pits 3 weeks postinfection (Figure S2A), the number of apoptotic cells in the pits was similar in both control and Hp-infected animals at 8 weeks postinfection (Figure 1B, left panels), indicating that Hp infection did not significantly affect gastric epithelial apoptosis in vivo. Then we orally administered Etp into the gerbil stomach 8 weeks after Hp infection, and apoptosis in the corpus and antrum tissues were investigated by TUNEL staining (Figure 1B, right panels). Etp administration increased the number of apoptotic epithelial cells in the pits infected with DcagA and DvirB7 as well as the noninfected controls; however, apoptosis in the superficial pit was significantly lower in the WT-infected animals (Figure 1B, right panels, and Figure 1C). Comparable results were obtained by staining for cleaved PARP, an apoptosis marker (Figures S2B–S2D). These data suggest that Hp can suppress apoptosis of gastric epithelial cells in vivo and that this bacterial activity is CagA dependent. CagA Inhibits Intrinsic Apoptosis via the MEK/ERK Pathway To evaluate whether Hp has antiapoptotic activity in vitro, gastric epithelial cells from Mongolian gerbils and AGS cells were left untreated or treated with U0126 or PD98059 (MEK inhibitors), and infected with WT, DcagA, DvirB7, DvirD4 (CagA secretion-deficient), DvirB11 (TFSSdeficient), or DvacA (VacA-deficient) derived from NCTC 11637, 26695, and ATCC 43579 strains. After incubating untreated cells with Etp or staurosporine (Stsp) to induce intrinsic apoptosis, the TUNEL- or annexin V-positive cell population was quantitated (Figures 2A and 2B). Though the extent of apoptosis varied among Hp strains, gerbil gastric epithelium and AGS cells infected with WT were more resistant to Etp/Stsp-induced apoptosis than the cells infected with DcagA, DvirB7, DvirD4, or DvirB11. However, when a similar experiment was performed in the presence of U0126/PD98059, WT infection did not prevent apoptosis, and the proportion of TUNEL- or annexin V-positive cells was similar among all Hp strains. Although Hp-infected epithelium not treated with Stsp or Etp could slightly induce apoptosis compared to epithelium treated with Etp (Figure S3A), and many studies have indicated that VacA can induce apoptosis (Boquet et al., 2003; Yamasaki et al., 2006), the apoptotic activity of DvacAinfected cells in these experimental conditions was similar to WT (Figure 2B). These results suggested that CagA plays an important role in modulating apoptosis. To determine whether the intrinsic or extrinsic apoptotic pathway is involved in the antiapoptotic activity caused by Hp infection, AGS cells were infected with Hp and incubated in the presence or absence of Stsp, Etp, Bak BH3 peptide (BH3) (intrinsic apoptosis inducers) or anti-Fas

Cell Host & Microbe 2, 250–263, October 2007 ª2007 Elsevier Inc. 251


Figure 1. Hp-CagA Prevents Apoptosis and Pit Loss in Mongolian Gerbils (A) Oral administration of Etp induces apoptosis in the gastric pits and exfoliation of pit layers. Animals were left untreated or given Etp orally, and 14 hr posttreatment, the gerbils were sacrificed and stomach sections were stained by the TUNEL method (apoptotic cells, green), Cy3-AAA (pit cells, red), Texas red-GS II (neck cells, purple), and DAPI (DNA, blue). The black and white arrowheads and yellow arrow, respectively, point to the pit surface, muscularis mucosae, and the exfoliated mucous surface layers containing apoptotic cells. Scale bar, 100 mm. (B and C) Hp-WT infection prevents Etp-induced apoptosis in pit cells. Animals (n = 8 per group) were each inoculated with Hp, and 8 weeks after infection, Etp was administered intragastrically. The gerbils were sacrificed, and stomach sections were immunostained by the TUNEL method (apoptotic cells, green), with anti-Hp antibody (blue), and with TOPRO-3 (DNA, red). The number of TUNELpositive and TO-PRO-3-positive cells in the gastric pits in an area approximately 350 mm wide was counted. More than ten areas per sample were measured (n = 80). All plots show the 25th to 75th percentiles (boxes) and 5th to 95th percentiles (whiskers). The line in each box represents the median. Scale bar, 100 mm.

antibody (extrinsic apoptosis inducer) (Figure 2C). The results showed that WT but not DcagA infection decreased the TUNEL-positive cell score when apoptosis was induced with Stsp, Etp, and BH3. This decrease in TUNELpositive AGS cells infected with WT was antagonized by PD98059 treatment. Because WT infection had less effect on apoptosis induced by anti-Fas antibody than on apoptosis induced by Stsp, Etp, or BH3 (Figure 2C), the intrinsic pathway appears to be the major pathway that stimulates CagA antiapoptotic activity. When apoptosis was induced by Stsp or Etp in WT-infected AGS cells, there were reduced levels of activated caspase-3/7 and -9 but not caspase-8 compared to DcagA- or DvirB7-infected AGS cells (Figure S3B). However, there were no appreciable differences in caspase levels between WT and DcagA during anti-Fas antibody-induced apoptosis (Figure S3B). Taken together, these results strongly suggested that

CagA has the ability to inhibit the intrinsic apoptotic pathway by activating MEK. GRB2, CRK, MEK, SRE, and SRF Are Intrinsic Host Elements that Induce the Antiapoptotic Activity of CagA To pursue if the CagA antiapoptotic pathway is MEK dependent, AGS cells overexpressing DsRed-tagged dominant-negative MEK (MEK-DN) (Figures 3A and 3B) were infected with WT and exposed to various intrinsic apoptosis inducers. In the cells expressing MEK-DN, the level of apoptosis was similar after WT and DcagA infection (Figure 3C), suggesting that CagA-dependent antiapoptosis requires MEK activation. Because CagA activates the MEK/ERK pathway by interacting with GRB2, CRK, or SHP2 via the CagA PY region, we investigated the involvement of these proteins in the CagA-directed



Figure 2. Hp Induces CagA-Dependent Inhibition of the Intrinsic Apoptosis Pathway via the MEK/ERK Pathway (A) Hp CagA antagonizes the apoptotic activity of Etp. Gastric epithelial cells from Mongolian gerbils were not infected () or infected with WT or the various isogenic mutant Hp strains, treated (+) or not treated () with U0126, and then exposed to Etp. The percentages of TUNEL-positive cells were calculated in arbitrary units set to a value of 100 for uninfected cells not treated with U0126. The results represent the average of three separate experiments (each n = 20). Data are means ± SD. (B) Hp CagA antagonizes the apoptotic activity of Stsp. AGS cells were not infected () or infected with WT or the various isogenic mutant strains of Hp, treated (+) or not treated () with PD98059, and then exposed to Stsp. The percentages of annexin V-positive cells are shown as fold increase over the noninfected cells not treated with PD98059. Data are means ± SD (n = 3). (C) The antiapoptotic activity of CagA is specific for the intrinsic pathway. AGS cells were not infected () or infected with Hp WT or DcagA, treated (+) or not treated () with PD98059, and then exposed to Stsp, Etp, BH3, or anti-Fas antibody and cycloheximide (Anti-Fas). The cells were fixed and stained by the TUNEL method. The percentages of TUNEL-positive cells were calculated after examination of at least 500 cells. The percentages of TUNEL-positive cells were calculated in arbitrary units set to a value of 100 for uninfected cells not treated with PD98059. The results represent the average of three separate experiments (each n = 20). Data are means ± SD.

antiapoptosis using siRNAs that specifically target GRB2, CRK, and SHP2 (Figure S4). Knocking down GRB2 and to a lesser extent CRK, but not SHP2, reduced the phosphorylation levels of MEK (Figures 3D and Figure S4C). Knockdown of GRB2 or CRK inhibited CagA-dependent antiapoptosis (Figure 3E), implying that CagA suppresses apoptosis via MEK/ERK activation. MEK/ERK activation induces phosphorylation of ternary complex factors (TCFs) components (Price et al., 1995), a subfamily of ETS domain transcription factors that bind and activate SREs in the promoters of target genes with the help of a second transcription factor, serum response factor (SRF). The SRF gene is also a target for TCFs/SRE, thereby providing a positive feedback loop in which TCF activation leads to enhanced expression of its partner protein SRF (Kasza et al., 2005). To determine whether infection by Hp activates the SRE or SRF promoter, AGS cells transfected with a reporter construct

containing these promoters (pSRE/pSRF-luc) were infected with Hp. As shown in Figure S5A, when the transfectants were infected with WT, but not with DcagA or DvirB7, the SRE level increased 2.7 fold, and SRF promoter activity increased 1.6 fold. These results were compatible with a previous report that showed similar findings using transient cagA transfectants (Hirata et al., 2002). When the AGS transfectants were pretreated with PD98059 or U0126 (MEK inhibitors), SRE activity and SRF activity in WT-infected cells decreased and were comparable to levels in DcagA or DvirB7-infected cells (Figure S5A). The fact that the levels of SRE, SRF, and NF-kB, but not of AP-1, were significantly lower in AGS cells expressing DPY (CagA lacking the PY region and unable to interact with GRB2 or CRK) than in AGS cells expressing CagA (Figure S5B) suggested that CagA is capable of stimulating SRE- and SRF-driven gene transcription via the GRB2/CRK-MEK pathway.



Figure 3. CagA-Dependent Antiapoptotic Activity Is Mediated via GRB2, CRK, and MEK (A) Western blot analysis of DsRed-MEK-DN-overexpressing AGS cells. AGS cells were transfected with DsRed (pDsRed-monomer) or DsRed-MEKDN (pDsRed-monomer-MEK-DN) expression vectors. Total cell lysates were separated by SDS-PAGE and subjected to western blot with the antibodies indicated. The white arrowheads point to nonspecific bands. (B) Overexpressed DsRed-MEK-DN does not affect mitochondrial morphology or cell shape. AGS cells transfected with DsRed-MEK-DN- or DsRed-expression vectors were fixed and stained with DAPI (DNA, blue) and anti-HSP70 (mitochondria, green). Scale bar, 20 mm. (C) CagA-induced antiapoptotic activity depends on MEK activation. AGS cells transfected with DsRed-MEK DN or DsRed-expression vectors were not infected () or infected with the noted Hp strains and then exposed to Stsp or Etp. The percentages of TUNEL-positive cells were calculated in arbitrary units set to a value of 100 for the noninfected cells not treated with U0126. The results represent the average of three separate experiments (each n = 20). Data are means ± SD. (D) Knockdown of GRB2 or CRK, but not SHP2, reduces CagA-induced MEK activation. AGS cells were transfected with siRNA oligonucleotides for luciferase (control: Luc), GRB2-1, -2, -3, CRK-1, -2, -3, or SHP2-1, -2, -3, and then not infected () or infected with Hp for 5 hr. Total cell lysates were separated by SDS-PAGE and subjected to western blot. The intensity of pMEK was calculated in arbitrary units set to a value of 1 for uninfected cells treated with luciferase siRNA. The results represent the average of three separate experiments. Data are means ± SD. (E) CagA-induced antiapoptotic activity is reduced by GRB2 or CRK knockdown. AGS cells were transfected with the indicated siRNA oligonucleotides, not infected () or infected with WT, DcagA, or DvirB7 Hp, and then treated with Etp. The percentages of TUNEL-positive cells were calculated in arbitrary units set to a value of 100 for uninfected cells treated with luciferase siRNA. The cells were fixed and stained by the TUNEL method. The percentages of TUNEL-positive cells were calculated after examining at least 1000 cells and are represented in arbitrary units set to a value of 100 for uninfected cells not treated with PD98059. The results represent the average of three separate experiments (each n = 20). Data are means ± SD.



CagA Increases MCL1 via SRE/SRF at Both the Transcriptional and Translational Levels Because activation of SRE/SRF-driven gene transcription affects levels of gene expression, including genes that inhibit the intrinsic pathway for apoptosis (Vickers et al., 2004), we quantitated mRNA levels of several BCL2 family proteins in Hp-infected AGS cells. Analysis of mRNA by RT-PCR showed that WT infection induced an approximately 1.5-fold increase in MCL1 mRNA level compared with DcagA or DvirB7 infection (Figure 4A). MCL1 protein levels in AGS cells were similarly increased by WT infection, approximately 1.6-fold greater than protein levels in AGS cells infected with the DcagA or DvirB7 mutants (Figure 4B). U0126 and BAY43-9006, MEK1 and RAF1 inhibitors, respectively, also blocked MCL1 production, whereas SB202190, SP600125, LY294002, and BAY117082, inhibitors of p38, JNK, PI3K, and NF-kB, respectively, did not block MCL1 production (Figure 4B). Because MCL1 is rapidly degraded by the ubiquitinproteasome pathway (Bannerman et al., 2001), we determined whether CagA is capable of modulating MCL1 ubiquitination by pretreating Hp-infected AGS cells with MG132 (proteasome inhibitor) to stabilize ubiquitinated MCL1. MG132 treatment of AGS cells did not alter the CagAdependent increase in MCL1 accumulation (Figure S6), indicating that the increase in MCL1 levels during WT infection was not due to modification of MCL1 ubiquitination. To confirm the role of CagA in stimulating MCL1 expression in Hp-infected gastric cells, we carried out transient transfection assays with a MCL1 promoter-driven luciferase reporter construct in AGS cells (pMCL1-WT-luc; Figure 4C). WT Hp infection efficiently increased the activity of the MCL1 promoter compared with the DcagA or DvirB7 mutants (Figures 4D and 4E). Introducing point mutations into the CArG (CC[A/T-rich]6GG) motif of SRE in pMCL1-WT-luc (pMCL1-CArG mut-luc) severely reduced the promoter activity in WT-infected cells (Figures 4C and 4E). Collectively, these findings demonstrate that CagA induces SRE/SRF-dependent transcriptional activity of the MCL1 gene and upregulates its expression. To further demonstrate that MCL1 upregulation by CagA is necessary for Hp’s antiapoptotic activity, we generated AGS cells in which MCL1 expression was decreased by siRNA (Figure S7). The results showed that WT-infected AGS cells treated with MCL1 siRNA were equally sensitive to Stsp-induced apoptosis as DcagA- or DvirB7-infected cells (Figure 5A). To show that the increase in MCL1 levels by CagA impacts the antiapoptotic activity, we created AGS cells transiently overexpressing DsRed-tagged MCL1 (Figures 5B and 5C). DsRed-MCL1 overexpressing cells infected with DcagA, DvirB7, or WT were resistant to Stsp- or Etp-induced apoptosis (Figure 5D), further demonstrating that the antiapoptotic activity caused by Hp infection results from increased MCL1 expression.

were inoculated with WT, DcagA, or DvirB7, and MCL1 mRNA expression in the stomach was measured by RTPCR 8 weeks later. MCL1 mRNA was increased more than 2.5-fold in WT-infected animals compared with the uninfected controls and animals infected with DcagA or DvirB7 (Figure S8A). MCL1 mRNA levels in Hp-infected animals were comparable to levels in Hp-infected gastric epithelium from Mongolian gerbils in vitro (Figure S8B), suggesting that the Hp mechanisms that induce MCL1 mRNA in the gastric epithelium in vitro are similar to the mechanisms in vivo. Immunohistochemical analysis with MCL1 antibody revealed thickened gastric pits and strong expression of MCL1 in both the corpus and antrum when infected with WT, but not with DcagA or DvirB7 (Figure S8C and Figures 6A and 6B). Consistent with the staining patterns of MCL1, increased levels of pERK were observed in the pit region in both the corpus and antrum when infected with WT, but not with DcagA or DvirB7 (Figures 6C and 6D). To assess whether the hyperplasia in the pits was attributable to pit cell proliferation, specimens were stained with antibodies to proliferating cell nuclear antigen (PCNA), AAA, or GSII lectin. When infected with WT, but not with DcagA or DvirB7, the numbers of PCNA-positive cells increased in areas around the isthmus and neck in the corpus of the stomach, whereas in the antrum of the stomach, PCNA-positive cells were confined to the isthmus region in the noninfected control and WT-, DcagA-, and DvirB7-infected animals (Figures 7A and 7B). Interestingly, PCNApositive cells were rarely detected in the pit region in the corpus and antrum in WT-, DcagA-, and DvirB7-infected animals and the noninfected controls. The immunohistochemical patterns of MCL1, pERK, and PCNA were similar in gerbils inoculated with WT or its isogenic mutants derived from the ATCC 43504 strain. However, at 8 weeks postinfection, WT ATCC 43504 showed more severe inflammation than WT NCTC 11637 and a marked increase in the number of PCNA-positive cells in the corpus/antrum transitional zone (data not shown). Based on the results, we speculated that the pit hyperplasia associated with WT infection is due not only to increased pit cell proliferation, but also to a decrease in abrasion of surface epithelium from the pits. As indicated by previous studies (Camorlinga-Ponce et al., 2004; Rieder et al., 2005), the density of Hp colonization in the gerbil stomach was significantly higher with WT than with DcagA or DvirB7 (Figure 7C). The differences in colonization densities between WT and mutant strains cannot be attributed to differences in the capacity to bind the gastric epithelium; there were no significant differences between WT and its isogenic mutant strains in the ability of Hp to bind the gastric epithelium in vitro (Figure S1B). These results further support the notion that CagA delivered by Hp into the pit epithelium plays a pivotal role in upregulating MCL1 expression, resulting in decreased apoptosis and pit loss and increased bacterial colonization (Figure 7D).

CagA-Induced MCL1 Expression in the Pit Epithelium Augments Hp Colonization To confirm that CagA suppresses apoptosis of gastric epithelium by upregulating MCL1 in vivo, Mongolian gerbils

DISCUSSION The gut epithelium constantly undergoes rapid self-renewal through the generation of epithelial progenitors at



Figure 4. CagA Induces MCL1 Transcription and Protein Expression via SRE/SRF (A) CagA upregulates the MCL1 mRNA level. AGS cells were not infected () or infected with the noted Hp strain for 4 hr. The expression level of the MCL1, BCL2, BCL2L1, and BAX genes was determined by real-time PCR. The results represent the average of four separate experiments (each n = 3). Data are means ± SD. (B) CagA upregulated the MCL1 protein expression level via RAF1 and MEK activation. AGS cells were not infected () or infected with the noted Hp strain and exposed or not exposed () to the inhibitors indicated for 4.5 hr. Total cell lysates were separated by SDS-PAGE and subjected to western blot with the noted antibodies. The intensity of MCL1 was normalized to the intensity of Actin and calculated in arbitrary units set to a value of 1 for uninfected cells not exposed to the inhibitor. The results represent the average of three separate experiments. Data are means ± SD. (C) Schematic representation of MCL1 reporter plasmids. The numbers of base pairs in the MCL1 50 -genomic flanking sequence are shown with the MCL1 transcription start site designated as +1. The mutated sequence in the SRE region of the construct (pMCL1-CArG mut-luc) is shown in red italics. (D) Hp-CagA induces MCL1 promoter activity. AGS cells transfected with MCL1 reporter plasmids were infected with Hp for 1.5, 3, 4.5, and 6 hr, and the lysates were measured by the luciferase assay. The luciferase activity is reported as the average fold induction compared with noninfected control from at least three independent experiments. Data are means ± SD (n = 4).



Figure 5. MCL1 Expression Is Indispensable for Hp-CagA-Induced Antiapoptotic Activity (A) CagA-induced antiapoptotic activity is reduced by MCL1 knockdown. AGS cells were transfected with siRNA oligonucleotides for the luciferase control (Luc), MCL1-1, -2, or -3, not infected () or infected with WT, DcagA, or DvirB7 Hp, and then treated with Etp. The percentages of TUNEL-positive cells were calculated after examining at least 1000 cells and are represented in arbitrary units set to a value of 100 for uninfected cells treated with luciferase siRNA. The results represent the average of three separate experiments (each n = 20). Data are means ± SD. (B) Western blot analysis of DsRed-MCL1expressing AGS cells. Total cell lysates from AGS cells transfected with DsRed (pDsRedexpress) or DsRed-MCL1 (pDsRed-MCL1) expression vectors were separated by SDSPAGE and subjected to western blot with the noted antibodies. Actin served as a loading control. (C) Overexpressed DsRed-MCL1 localizes mainly to the mitochondria and does not affect mitochondrial morphology or cell shape. AGS cells transfected with DsRed-MCL1 or DsRed expression vectors were fixed and immunostained with anti-HSP70 antibody (mitochondria, green) and DAPI (DNA, blue). Scale bar, 20 mm. (D) MCL1 overexpression compensates for CagA-dependent antiapoptotic activity. AGS cells transfected with DsRed or DsRed-MCL1 expression vectors were infected with Hp and then treated with Stsp or Etp. The percentages of TUNEL-positive cells were calculated after examining at least 1000 cells and are represented in arbitrary units set to a value of 100 for uninfected cells treated with luciferase siRNA. The results represent the average of three separate experiments (each n = 20). Data are means ± SD.

the isthmus of gastric crypts and cell death at superficial pit cells. The rapid turnover of epithelial cells serves as an intrinsic defense system against microbial infection, and disruption of this physiological equilibrium by suppressing apoptosis has serious consequences for gastric illness. In the present study we induced apoptosis in a Mongolian gerbil stomach model by orally inoculating Etp, an intrinsic apoptosis inducer, in order to evaluate how Hp infection affected apoptosis of gastric epithelial cells. Using this model, we have provided convincing in vivo and in vitro evidence that CagA delivered by the Hp TFSS into the gastric epithelium suppressed apoptosis by upregulating MCL1 expression. Importantly, this bacterial activity greatly contributed to bacterial colonization of the gut epithelium, where CagA/MCL1 interplay played a pivotal role (Figures 1, 6, and 7). CagA can induce proliferation of gastric epithelial cells in vitro by interacting with host factors such as GRB2 and CRK, where the stimulation of MEK/ERK pathway is important (see Introduction). As gastric epithelial cells rapidly turn over, Hp can pro-

mote colonization by enhancing antiapoptosis, a property that is an important bacterial strategy to sustain chronic infection. Several reports have indicated that CagA activates signaling pathways other than the MEK/ERK pathway, including p38, JNK, PI3K/AKT, and NF-kB pathways (Brandt et al., 2005) (Figure S5B). Furthermore, the Hp peptidoglycan is also thought to activate NF-kB (Viala et al., 2004). CagA also contributes to modulation of other apoptosis-regulating factors such as c-IAP-2, BCL2, or Bad phosphorylation in combination with some signaling pathways (Brandt et al., 2005; Choi et al., 2003; Yanai et al., 2003; Zhu et al., 2007). Although the importance of each CagA-induced signaling pathway on cell death is still unclear, we speculate that balance between cell death and survival in the gastric mucosa is orchestrated by multiple molecular mechanisms that create an optimum environment for bacterial colonization. Microarray analysis previously indicated that MCL1 is one of the host genes upregulated by Hp infection in

(E) Activation of the MCL1 reporter gene by Hp-CagA depends on its SRE region. AGS cells transfected with MCL1 reporter plasmids were infected with Hp for 4.5 hr, and the lysates were measured by the luciferase assay. The luciferase activity is reported as the average fold induction compared with the noninfected control from at least three independent experiments. Data are means ± SD (n = 4).



Figure 6. Hp-CagA Induces MCL1 and pERK Expression in the Gastric Pits of Hp-Infected Mongolian Gerbils (A and B) MCL1 protein expression is upregulated in the gastric pit of Hp WT-infected Mongolian gerbils. The gastric sections from Hp (NCTC 11637)-infected animals (n = 8 per group) were immunostained with anti-MCL1 antibody (green) and Cy3-conjugated AAA (red). (A) Neck cells were labeled with Texasred-GS II (blue). Scale bar, 100 mm. (B) DNA was labeled with TO-PRO-3. Fluorescence intensity within an area of approximately 100 mm high (from the top of pits to neck region) and approximately 350 mm wide was assessed by confocal microscopy. The intensity of FITClabeled MCL1 was calculated as a ratio to that of TO-PRO-3. More than ten areas per sample were measured (n = 80) Data are means ± SD. (C and D) Number of pERK-positive cell increases in the gastric pit of Hp WT-infected Mongolian gerbils. The gastric sections from Hp (NCTC 11637)-infected animals (n = 8 per group) were immunostained with anti-pERK antibody (green) and Cy3-conjugated AAA (red). (C) Neck cells were labeled with Texasred-GS II (blue). Scale bar, 100 mm. (D) DNA was labeled with DAPI (blue). Fluorescence intensity within an area of approximately 100 mm high (from the top of pits to neck region) and approximately 350 mm wide was assessed by confocal microscopy. The intensity of FITClabeled pERK was calculated as a ratio to that of DAPI. More than three areas per sample were measured (n = 24). Data are means ± SD.

a CagA-dependent manner (El-Etr et al., 2004). The upstream region of the MCL1 gene has several putative consensus binding sites for transcription factors, including E2F1 (Croxton et al., 2002), PI3K/AKT-activated STAT3 (Isomoto et al., 2005), CREB (Wang et al., 1999), hypoxia-induced HIF-1 (Piret et al., 2005), ERK-activated SRF, Elk-1, and Sp1 (Townsend et al., 1999). Phosphorylation of MCL1 induced by GSK-3 in a B cell line can be inactivated by PI3K/AKT, and phosphorylated MCL1 is ubiquitinated and degraded by proteasomes (Maurer et al., 2006). Although the PI3K-dependent regulation mechanism seems to be conspicuous in B cells, MCL1 upregulation by CagA in gastric epithelial cells was unaffected by PI3K activation (Figure 4B). Instead, we showed that CagA-mediated stimulation of RAF1/MEK/ERK is indispensable for activation of the MCL1 promoter.

Some carbohydrate structures such as Lewis antigens on surface gastric cells act as ligands for Hp adhesion (Ilver et al., 1998; Mahdavi et al., 2002). By contrast, it has been proposed that O-glycans with a terminal a1,4-linked N-acetylglucosamine in deeper portions of the gastric mucosa have antibiotic functions against Hp (Kawakubo et al., 2004), indicating that the majority of direct interactions between Hp and the infected host take place at the superficial pit epithelium of the stomach. In the gerbil stomach Hp model, the antiapoptotic effects conferred by CagA/MCL1 on cells resulted in hyperplasia in the pits. Consistent with the previous report that hyperplasia in the pits is favorable for Hp expansion and colonization (Fox et al., 1999), WT colonization in the gerbil stomach was considerably higher than cagA-defective mutants (Figure 7C). Taken together, these data suggest that, by



Figure 7. Hp-CagA Prevents Apoptotic Loss of Pit Cells and Promotes Bacterial Colonization in Mongolian Gerbils (A and B) Hyperplasia in the pits is not attributable to proliferation of pit cells. At 8 weeks after infection, the gerbils were sacrificed, and stomach sections were immunostained with PCNA (proliferating cells, green), AAA (pit cells, red), and GS II (neck cells, blue). The isthmus region is the area containing a large number of PCNA-positive cells between the pit region and neck region. (A) Scale bar, 100 mm. (B) The number of PCNA-positive cells in an area approximately 100 mm high from the top of pits to the muscularis mucosae and approximately 350 mm wide was counted by confocal microscopy. More than ten areas per sample were measured (n = 80). All plots show the 25th to 75th percentiles (boxes) and 5th to 95th percentiles (whiskers). The line in the box represents the median. (C) Hp-WT efficiently colonize the stomach of Mongolian gerbil. Animals (n = 8 per group) were inoculated with Hp. At 8 weeks after infection, animals were sacrificed, and a quantitative culture assay was performed on gastric specimens. All plots show the 25th to 75th percentiles (boxes) and 5th to 95th percentiles (whiskers). The line in each box represents the median (n = 8). (D) Hp CagA upregulates MCL1 expression and protects gastric epithelial cells from the intrinsic pathway of apoptosis.

injecting CagA into the epithelial cells, Hp makes the microenvironment in the pits more suitable for long-term colonization. In Hp-infected gerbils, enhanced apoptosis levels were similar between WT and isogenic mutant strains, including DcagA and DvirB7, 3 weeks post-Hp inoculation (Figures S2A and S2C), but no significant apoptosis was observed 8 weeks postinfection (Figure 1B and Figure S2D). These data are consistent with a previous chronological study using a Mongolian gerbil animal model (Peek et al., 2000). This report along with our study indicate that Hp infection increases apoptosis to maximum levels at 2–4 weeks postchallenge, with no differences in apoptotic indices between WT, DcagA, and DvacA, followed by a decrease in apoptotic indices to baseline. Though Hp induced MCL1 production in infected epithelium within a few hours of infection in vitro (Figure 4 and Figure S8B), no inhibitory effect of apoptosis could be seen in the

Hp-infected gerbil’s stomach 3 weeks postinfection in vivo (Figure S2). Since we could not detect Hp in the stomach 3 weeks postinfection (data not shown), one reason for delayed antiapoptotic effect in vivo would be the gastric mucus layer, which might reduce the intimate attachment of Hp to epithelium surface in vivo. There are many reports on apoptosis induction in Hp infection both in vitro and in vivo. Our data indicate that CagA and TFSS machineries are not among the bacterial factors that induce apoptosis in an Hp-infected gerbil stomach in vivo or gastric epithelium in vitro. Rudi et al. reported that the CD95 (Fas) receptor and its ligand, which are activated by an unidentified soluble Hp fraction, are involved in Hp-induced gastric epithelium apoptosis in human biopsies and in vitro (Rudi et al., 1998). Thus, it is speculated that CD95-mediated apoptosis might be included in the apoptosis induced during the early phase of Hp infection.



In contrast, proliferation rates of gastric cells in the lamina propria were higher at 8 weeks (Figure 7A) than at 3 weeks post-WT Hp inoculation (data not shown). Moreover, we found that, 8 weeks after Hp ATCC 43504 WT infection, the corpus/antral transitional zone showed inflammation with severe hyperplasia and several lymphoid follicles (data not shown). Thus, our data are fully compatible with other studies reporting that later in infection the gastric mucosa shifts from an apoptotic response to a CagA-dependent hyperproliferative response, which may heighten the risk of developing gastric neoplasia (Peek et al., 1997), even though the degree of inflammation and hyperproliferation may differ between each WT Hp strain. In our studies, greater MCL1 mRNA/protein upregulation and increased antiapoptotic effects on the epithelium were seen in Hp-infected gerbils in vivo (Figures S8A and S2D and Figures 1B and 6A) than in vitro using AGS cells (Figures 2C, 4A, and 4B). These differences between in vivo and in vitro data may be attributed to the general ability of cancer cell lines, including AGS, to resist apoptosis. Indeed, primary cultures of gastric epithelium from gerbils had higher sensitivity to Etp (Figure S3A) (data not shown) and higher rates of CagA-dependent MCL1 upregulation (Figure S8B) than AGS cells (Figure 4A). Although molecular studies with Mongolian gerbils are still limited because of the lack of genomic information, Hp infection and pathogenesis in Mongolian gerbils generally reflect human infection and are similar to other animal models. Intriguingly, immunohistochemical analysis of the stomach of Hp-infected gerbils showed little Hp in the isthmus to neck region, the deeper portion of the gastric mucosa, where we observed a CagA-dependent increase in proliferating cells (Figure 7A), and a region of the stomach that occasionally develops gastric cancer and MALT lymphoma (Houghton and Wang, 2005). However, CagAinduced ERK activation and antiapoptotic action mostly occurred in the superficial pit cells (Figures 1A, 1B, 6A, and 6C), where the vast majority of bacteria had colonized (Figure 1C). Although we could not rule out the possibility that a small number of Hp colonized deeper in the gastric crypts and stimulated proliferation of stem cells, direct contact between multiplying Hp and superficial gastric epithelium is more likely the major route of CagA delivery and induction of pit hyperplasia. Alternatively, Hp-induced inflammatory mediators, such as mediators derived from epithelial cells, activated leukocytes, or migratory immune cells may also stimulate proliferation of epithelial progenitors residing deeper in the crypts. For example, a recent study showed that the gastric adenocarcinoma induced by chronic infection of mice with Helicobacter felis originates from circulating bone marrow-derived cells, not from resident gastric cells (Houghton et al., 2004). Moreover, a recent study indicates that persistent Hp colonization in the stomach and continuous Hp uptake through intestinal Peyer’s patches are prerequisites for inducing inflammation in the stomach (Nagai et al., 2007). Therefore, it is reasonable to speculate that long-term Hp

colonization of the gastric epithelium increases the risk of gastric carcinoma or MALT lymphoma. In summary, we have obtained evidence that CagA delivered by Hp into the gastric epithelium plays a pivotal role in preventing apoptosis and suppressing the rapid turnover of superficial pit cells. By pharmacologically inducing apoptosis in Mongolian gerbils and in AGS cells, we have demonstrated that Hp infection of gastric epithelium is capable of increasing MCL1 expression via the MEK/ERK/SRE pathway to enhance cell survival and Hp colonization. EXPERIMENTAL PROCEDURES Strains and Cell Culture Hp strain NCTC 11637 and its isogenic mutants DcagA, DvirD7, and DvacA; strain 26695, DvirD4, and DvirD7; and strain ATCC 43579, DcagA, and DvirD11 have been described previously (Asahi et al., 2000; Mimuro et al., 2002; Suzuki et al., 2005). Isogenic cagA and virB7 null mutants derived from ATCC 43504 were constructed by insertional mutagenesis, using aphA (conferring kanamycin resistance). All strains were cultured according to standard procedures (Mimuro et al., 2002). AGS cells were maintained in Dulbecco’s modified Eagle medium/F-12 (Ham’s) (Invitrogen) containing 10% fetal bovine serum (FBS), and NCI-N87 cells were maintained in RPMI-1640 (SigmaAldrich) containing 10% FBS. Primary gastric epithelial cells were prepared as described previously (Viala et al., 2004). Bacterial Infection AGS cells were infected with Hp at a multiplicity of infection of 50 per cell. In the experiments using inhibitors, AGS cells were incubated with 50 mM PD98059, 50 mM U0126, 20 mM SB202190, 50 mM SP600125, 50 mM LY294002, 5 mM BAY43-9006, 2 mM BAY11-7082, or 40 mM MG132 for 30 min following Hp infection. For western blot analysis, equal amounts of protein from each sample were separated by SDSPAGE and transferred to a nitrocellulose membrane. To compare protein levels, bands were quantified by densitometry, analyzed with ImageJ software, and normalized with the protein level of actin (Figure 4B and Figure S6) or MEK (Figure 3D). Transient Expression in Cultured Cells Transfection was carried out with FuGENE6 Transfection Reagent (Roche). At 18 hr after transfection, the cells were subjected to Hp infection, immunofluorescence staining, or cell lysis. Apoptosis Assays AGS cells infected with Hp for 3 hr were exposed to 1 mM Stsp (SigmaAldrich), 300 mM Etp (Sigma-Aldrich), 50 mM Bak BH3 peptide (Calbiochem), or 100 ng/ml Anti-Fas antibody with 2 mg/ml of cycloheximide for 4 hr. For annexin V staining, cells were harvested and stained with annexin V-FLUOS (Roche) according to the manufacturer’s instructions. Flow cytometric analysis was performed with a FACSCalibur flow cytometer (BD Biosciences) and CellQuest and FlowJo softwares. The fluorescence intensity level of unstained cells was calibrated as negative. A TUNEL assay was performed with the DeadEnd fluorometric TUNEL system (Promega). The cells were analyzed with a microscope equipped with a cooled CCD camera and with IPLab Spectrum software (Signal Analytics Corp). Caspase-9, -8, and -3/7 activity was analyzed using caspase-Glo assays (Promega). Hp Gastric Infection of Mongolian Gerbils Six-week-old male MGS/Sea Mongolian gerbils (Meriones unguiculatus; CLEA Japan Inc.) fasted for 20 hr were intragastrically inoculated with a bacterial culture containing 109 cfu of Hp in Brucella broth containing 5% (v/v) of FCS twice daily for 2 consecutive days. Control animals were orally administered Brucella broth containing 5% (v/v) FCS. After 8 weeks, the experimental animals were given 30 mg/5 ml/g



body weight of Etp dissolved in 20% DMSO-80% saline (v/v) orally. Control animals were given 20% DMSO-80% saline (v/v). After 14 hr, the stomach of each infected Mongolian gerbil was opened along the greater curvature. To quantitatively isolate Hp, the stomach was excised, weighed, and homogenized. Serial dilutions were plated on Hp-selective agar plates (Eiken Chemical Co.) and incubated under microaerophilic conditions at 37 C for 4 days, after which the cfu were counted. Colonization data points of 1 3 103 cfu were the minimal detection limit of the assay. For RNA isolation, the tissue was immediately frozen in liquid nitrogen. Sections for histological analysis were fixed in 4% paraformaldehyde in PBS. The gastric tissue embedded in tissuefreezing medium (Jung Tissue Freezing Medium; Leica Microsystems) was frozen in liquid nitrogen and sectioned with a Leica cryostat (model CM1900). All animal experiments were conducted in accordance with the guidelines of the University of Tokyo for the care and use of laboratory animals and were approved by the ethics committee for animal experiments of the University of Tokyo. Immunofluorescence Microscopy Anguilla anguilla agglutinin (AAA; Sigma-Aldrich) was Cy3 labeled using Cy3 monofunctional reactive dye (Amersham Bioscience). Immunofluorescence staining was performed as described previously (Mimuro et al., 2002). For PCNA staining, acetone-fixed sections were stained with anti-PCNA antibody after epitope retrieval by heating in citrate buffer (10 mM sodium citrate buffer [pH 6.0]). For cleaved-PARP staining, acetone-fixed sections were stained with anti-PARP 214/215 cleavage site-specific antibody after epitope retrieval by heating in Tris-EDTA buffer (10 mM Tris buffer, 0.1 mM EDTA, 0.05% Tween 20 [pH 9.0]), followed by the tyramide amplification method (TSA-Plus Fluorescein System; Perkin Elmer Life sciences, MA). FITC-, TRITC-, or Cy5-conjugated secondary antibody was used to visualize rabbit or mouse antibodies, TO-PRO-3 (Molecular Probes) or DAPI (Sigma-Aldrich) was used to visualize DNA, and anti-HSP70 antibody was used to visualize mitochondria. The pERK signal was detected by using immunoreaction enhancer solution (Can Get Signal immunostain, Toyobo). Texas red-labeled Griffonia simplicifolia II (GSII) lectin (EY Laboratories) was used to visualize neck cells in stomach specimens, and Cy3-labeled AAA was used to visualize pit cells. The stained specimens were examined with a confocal laser-scanning microscope (LSM510; Carl Zeiss). Fluorescence intensity was measured with LSM510 version 3.2 software (Carl Zeiss). Statistical Analysis Differences between groups in the in vitro experiments were assessed by the paired, two-tailed Student’s t test. The data in Figures 1, 6, and 7 and Figures S2 and S8A were statistically analyzed by using the MannWhitney U test for unpaired groups. Supplemental Data The Supplemental Data include Supplemental Experimental Procedures and eight supplemental figures and can be found with this article online at http://www.cellhostandmicrobe.com/cgi/content/full/2/4/ 250/DC1/. ACKNOWLEDGMENTS We thank Tomoko Furukawa, Yoshiko Kawahara, and Melanie Hallard for technical help; Akira Nishizono, Toshio Fujioka, Richard M. Peek, and Steffen Backert for providing Hp strains; Osamu Igarashi, Kazunari Murakami, Takashi Nonaka, and Timothy Cragin Wang for helpful advice; and all members of the Sasakawa laboratory for helpful discussions and technical advice. This work was supported by a Grant-inAid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology of Japan (18689013 and 17659127 to H.M.); Japan Science and Technology Agency (to C.S.); the National BioResource Project in Japan (http://www.nbrp.jp/); and the Deutsche Forschungsgemeinschaft (SFB576, Project B1 to R.H.).

Received: January 4, 2007 Revised: August 11, 2007 Accepted: September 14, 2007 Published: October 10, 2007 REFERENCES Asahi, M., Azuma, T., Ito, S., Ito, Y., Suto, H., Nagai, Y., Tsubokawa, M., Tohyama, Y., Maeda, S., Omata, M., et al. (2000). Helicobacter pylori CagA protein can be tyrosine phosphorylated in gastric epithelial cells. J. Exp. Med. 191, 593–602. Backert, S., and Meyer, T.F. (2006). Type IV secretion systems and their effectors in bacterial pathogenesis. Curr. Opin. Microbiol. 9, 207–217. Bannerman, D.D., Tupper, J.C., Ricketts, W.A., Bennett, C.F., Winn, R.K., and Harlan, J.M. (2001). A constitutive cytoprotective pathway protects endothelial cells from lipopolysaccharide-induced apoptosis. J. Biol. Chem. 276, 14924–14932. Blaser, M.J. (1990). Helicobacter pylori and the pathogenesis of gastroduodenal inflammation. J. Infect. Dis. 161, 626–633. Blaser, M.J., Perez-Perez, G.I., Kleanthous, H., Cover, T.L., Peek, J.R.M., Chyou, P.H., Stemmermann, G.N., and Nomura, A. (1995). Infection with Helicobacter pylori strains possessing cagA is associated with an increased risk of developing adenocarcinoma of the stomach. Cancer Res. 55, 2111–2115. Boquet, P., Ricci, V., Galmiche, A., and Gauthier, N.C. (2003). Gastric cell apoptosis and H. pylori: Has the main function of VacA finally been identified? Trends Microbiol. 11, 410–413. Boucher, M.J., Morisset, J., Vachon, P.H., Reed, J.C., Laine, J., and Rivard, N. (2000). MEK/ERK signaling pathway regulates the expression of Bcl-2, Bcl-X(L), and Mcl-1 and promotes survival of human pancreatic cancer cells. J. Cell. Biochem. 79, 355–369. Brandt, S., Kwok, T., Hartig, R., Konig, W., and Backert, S. (2005). NF-kappaB activation and potentiation of proinflammatory responses by the Helicobacter pylori CagA protein. Proc. Natl. Acad. Sci. USA 102, 9300–9305. Budd, R.C., Yeh, W.C., and Tschopp, J. (2006). cFLIP regulation of lymphocyte activation and development. Nat. Rev. Immunol. 6, 196– 204. Camorlinga-Ponce, M., Romo, C., Gonzalez-Valencia, G., Munoz, O., and Torres, J. (2004). Topographical localisation of cagA positive and cagA negative Helicobacter pylori strains in the gastric mucosa; an in situ hybridisation study. J. Clin. Pathol. 57, 822–828. Choi, I.J., Kim, J.S., Kim, J.M., Jung, H.C., and Song, I.S. (2003). Effect of inhibition of extracellular signal-regulated kinase 1 and 2 pathway on apoptosis and bcl-2 expression in Helicobacter pylori-infected AGS cells. Infect. Immun. 71, 830–837. Croxton, R., Ma, Y., Song, L., Haura, E.B., and Cress, W.D. (2002). Direct repression of the Mcl-1 promoter by E2F1. Oncogene 21, 1359–1369. Eck, M., Schmausser, B., Haas, R., Greiner, A., Czub, S., and MullerHermelink, H.K. (1997). MALT-type lymphoma of the stomach is associated with Helicobacter pylori strains expressing the CagA protein. Gastroenterology 112, 1482–1486. El-Etr, S.H., Mueller, A., Tompkins, L.S., Falkow, S., and Merrell, D.S. (2004). Phosphorylation-independent effects of CagA during interaction between Helicobacter pylori and T84 polarized monolayers. J. Infect. Dis. 190, 1516–1523. Falk, P., Roth, K.A., Boren, T., Westblom, T.U., Gordon, J.I., and Normark, S. (1993). An in vitro adherence assay reveals that Helicobacter pylori exhibits cell lineage-specific tropism in the human gastric epithelium. Proc. Natl. Acad. Sci. USA 90, 2035–2039. Fox, J.G., Dangler, C.A., Taylor, N.S., King, A., Koh, T.J., and Wang, T.C. (1999). High-salt diet induces gastric epithelial hyperplasia and



parietal cell loss, and enhances Helicobacter pylori colonization in C57BL/6 mice. Cancer Res. 59, 4823–4828. Franco, A.T., Israel, D.A., Washington, M.K., Krishna, U., Fox, J.G., Rogers, A.B., Neish, A.S., Collier-Hyams, L., Perez-Perez, G.I., Hatakeyama, M., et al. (2005). Activation of beta-catenin by carcinogenic Helicobacter pylori. Proc. Natl. Acad. Sci. USA 102, 10646–10651. Galmiche, A., Rassow, J., Doye, A., Cagnol, S., Chambard, J.C., Contamin, S., de Thillot, V., Just, I., Ricci, V., Solcia, E., et al. (2000). The N-terminal 34 kDa fragment of Helicobacter pylori vacuolating cytotoxin targets mitochondria and induces cytochrome c release. EMBO J. 19, 6361–6370. Higashi, H., Tsutsumi, R., Muto, S., Sugiyama, T., Azuma, T., Asaka, M., and Hatakeyama, M. (2002). SHP-2 tyrosine phosphatase as an intracellular target of Helicobacter pylori CagA protein. Science 295, 683–686. Hirata, Y., Maeda, S., Mitsuno, Y., Tateishi, K., Yanai, A., Akanuma, M., Yoshida, H., Kawabe, T., Shiratori, Y., and Omata, M. (2002). Helicobacter pylori CagA protein activates serum response element-driven transcription independently of tyrosine phosphorylation. Gastroenterology 123, 1962–1971. Houghton, J., Stoicov, C., Nomura, S., Rogers, A.B., Carlson, J., Li, H., Cai, X., Fox, J.G., Goldenring, J.R., and Wang, T.C. (2004). Gastric cancer originating from bone marrow-derived cells. Science 306, 1568–1571.

Mills, J.C., Andersson, N., Hong, C.V., Stappenbeck, T.S., and Gordon, J.I. (2002). Molecular characterization of mouse gastric epithelial progenitor cells. Proc. Natl. Acad. Sci. USA 99, 14819–14824. Mimuro, H., Suzuki, T., Tanaka, J., Asahi, M., Haas, R., and Sasakawa, C. (2002). Grb2 is a key mediator of Helicobacter pylori CagA protein activities. Mol. Cell 10, 745–755. Nagai, S., Mimuro, H., Yamada, T., Baba, Y., Moro, K., Nochi, T., Kiyono, H., Suzuki, T., Sasakawa, C., and Koyasu, S. (2007). Role of Peyer’s patches in the induction of Helicobacter pylori-induced gastritis. Proc. Natl. Acad. Sci. USA 104, 8971–8976. Odenbreit, S., Puls, J., Sedlmaier, B., Gerland, E., Fischer, W., and Haas, R. (2000). Translocation of Helicobacter pylori CagA into gastric epithelial cells by type IV secretion. Science 287, 1497–1500. Oh, J.D., Karam, S.M., and Gordon, J.I. (2005). Intracellular Helicobacter pylori in gastric epithelial progenitors. Proc. Natl. Acad. Sci. USA 102, 5186–5191. Opferman, J.T., Iwasaki, H., Ong, C.C., Suh, H., Mizuno, S., Akashi, K., and Korsmeyer, S.J. (2005). Obligate role of anti-apoptotic MCL-1 in the survival of hematopoietic stem cells. Science 307, 1101–1104. Parsonnet, J., Friedman, G.D., Vandersteen, D.P., Chang, Y., Vogelman, J.H., Orentreich, N., and Sibley, R.K. (1991). Helicobacter pylori infection and the risk of gastric carcinoma. N. Engl. J. Med. 325, 1127–1131.

Houghton, J., and Wang, T.C. (2005). Helicobacter pylori and gastric cancer: A new paradigm for inflammation-associated epithelial cancers. Gastroenterology 128, 1567–1578.

Parsonnet, J., Hansen, S., Rodriguez, L., Gelb, A.B., Warnke, R.A., Jellum, E., Orentreich, N., Vogelman, J.H., and Friedman, G.D. (1994). Helicobacter pylori infection and gastric lymphoma. N. Engl. J. Med. 330, 1267–1271.

Ilver, D., Arnqvist, A., Ogren, J., Frick, I.M., Kersulyte, D., Incecik, E.T., Berg, D.E., Covacci, A., Engstrand, L., and Boren, T. (1998). Helicobacter pylori adhesin binding fucosylated histo-blood group antigens revealed by retagging. Science 279, 373–377.

Peek, R.M., Jr., Moss, S.F., Tham, K.T., Perez-Perez, G.I., Wang, S., Miller, G.G., Atherton, J.C., Holt, P.R., and Blaser, M.J. (1997). Helicobacter pylori cagA+ strains and dissociation of gastric epithelial cell proliferation from apoptosis. J. Natl. Cancer Inst. 89, 863–868.

Isomoto, H., Kobayashi, S., Werneburg, N.W., Bronk, S.F., Guicciardi, M.E., Frank, D.A., and Gores, G.J. (2005). Interleukin 6 upregulates myeloid cell leukemia-1 expression through a STAT3 pathway in cholangiocarcinoma cells. Hepatology 42, 1329–1338.

Peek, R.M., Jr., Wirth, H.P., Moss, S.F., Yang, M., Abdalla, A.M., Tham, K.T., Zhang, T., Tang, L.H., Modlin, I.M., and Blaser, M.J. (2000). Helicobacter pylori alters gastric epithelial cell cycle events and gastrin secretion in Mongolian gerbils. Gastroenterology 118, 48–59.

Jiang, X., and Wang, X. (2004). Cytochrome C-mediated apoptosis. Annu. Rev. Biochem. 73, 87–106.

Peek, R.M., Jr., and Blaser, M.J. (2002). Helicobacter pylori and gastrointestinal tract adenocarcinomas. Nat. Rev. Cancer 2, 28–37.

Karam, S.M., and Leblond, C.P. (1993). Dynamics of epithelial cells in the corpus of the mouse stomach. II. Outward migration of pit cells. Anat. Rec. 236, 280–296.

Piret, J.P., Minet, E., Cosse, J.P., Ninane, N., Debacq, C., Raes, M., and Michiels, C. (2005). Hypoxia-inducible factor-1-dependent overexpression of myeloid cell factor-1 protects hypoxic cells against tert-butyl hydroperoxide-induced apoptosis. J. Biol. Chem. 280, 9336–9344.

Kasza, A., O’Donnell, A., Gascoigne, K., Zeef, L.A., Hayes, A., and Sharrocks, A.D. (2005). The ETS domain transcription factor Elk-1 regulates the expression of its partner protein, SRF. J. Biol. Chem. 280, 1149–1155. Kawahara, T., Kuwano, Y., Teshima-Kondo, S., Sugiyama, T., Kawai, T., Nikawa, T., Kishi, K., and Rokutan, K. (2001). Helicobacter pylori lipopolysaccharide from type I, but not type II strains, stimulates apoptosis of cultured gastric mucosal cells. J. Med. Invest. 48, 167–174. Kawakubo, M., Ito, Y., Okimura, Y., Kobayashi, M., Sakura, K., Kasama, S., Fukuda, M.N., Fukuda, M., Katsuyama, T., and Nakayama, J. (2004). Natural antibiotic function of a human gastric mucin against Helicobacter pylori infection. Science 305, 1003–1006. Kuipers, E.J., Perez-Perez, G.I., Meuwissen, S.G., and Blaser, M.J. (1995). Helicobacter pylori and atrophic gastritis: Importance of the cagA status. J. Natl. Cancer Inst. 87, 1777–1780. Mahdavi, J., Sonden, B., Hurtig, M., Olfat, F.O., Forsberg, L., Roche, N., Angstrom, J., Larsson, T., Teneberg, S., Karlsson, K.A., et al. (2002). Helicobacter pylori SabA adhesin in persistent infection and chronic inflammation. Science 297, 573–578. Maurer, U., Charvet, C., Wagman, A.S., Dejardin, E., and Green, D.R. (2006). Glycogen synthase kinase-3 regulates mitochondrial outer membrane permeabilization and apoptosis by destabilization of MCL-1. Mol. Cell 21, 749–760.

Price, M.A., Rogers, A.E., and Treisman, R. (1995). Comparative analysis of the ternary complex factors Elk-1, SAP-1a and SAP-2 (ERP/ NET). EMBO J. 14, 2589–2601. Rieder, G., Merchant, J.L., and Haas, R. (2005). Helicobacter pylori cag-type IV secretion system facilitates corpus colonization to induce precancerous conditions in Mongolian gerbils. Gastroenterology 128, 1229–1242. Rudi, J., Kuck, D., Strand, S., von Herbay, A., Mariani, S.M., Krammer, P.H., Galle, P.R., and Stremmel, W. (1998). Involvement of the CD95 (APO-1/Fas) receptor and ligand system in Helicobacter pyloriinduced gastric epithelial apoptosis. J. Clin. Invest. 102, 1506–1514. Segal, E.D., Cha, J., Lo, J., Falkow, S., and Tompkins, L.S. (1999). Altered states: Involvement of phosphorylated CagA in the induction of host cellular growth changes by Helicobacter pylori. Proc. Natl. Acad. Sci. USA 96, 14559–14564. Selbach, M., Moese, S., Hauck, C.R., Meyer, T.F., and Backert, S. (2002). Src is the kinase of the Helicobacter pylori CagA protein in vitro and in vivo. J. Biol. Chem. 277, 6775–6778. Shibayama, K., Kamachi, K., Nagata, N., Yagi, T., Nada, T., Doi, Y., Shibata, N., Yokoyama, K., Yamane, K., Kato, H., et al. (2003). A novel apoptosis-inducing protein from Helicobacter pylori. Mol. Microbiol. 47, 443–451.



Strasser, A. (2005). The role of BH3-only proteins in the immune system. Nat. Rev. Immunol. 5, 189–200. Suzuki, M., Mimuro, H., Suzuki, T., Park, M., Yamamoto, T., and Sasakawa, C. (2005). Interaction of CagA with Crk plays an important role in Helicobacter pylori-induced loss of gastric epithelial cell adhesion. J. Exp. Med. 202, 1235–1247. Townsend, K.J., Zhou, P., Qian, L., Bieszczad, C.K., Lowrey, C.H., Yen, A., and Craig, R.W. (1999). Regulation of MCL1 through a serum response factor/Elk-1-mediated mechanism links expression of a viability-promoting member of the BCL2 family to the induction of hematopoietic cell differentiation. J. Biol. Chem. 274, 1801–1813. Viala, J., Chaput, C., Boneca, I.G., Cardona, A., Girardin, S.E., Moran, A.P., Athman, R., Memet, S., Huerre, M.R., Coyle, A.J., et al. (2004). Nod1 responds to peptidoglycan delivered by the Helicobacter pylori cag pathogenicity island. Nat. Immunol. 5, 1166–1174. Vickers, E.R., Kasza, A., Kurnaz, I.A., Seifert, A., Zeef, L.A., O’Donnell, A., Hayes, A., and Sharrocks, A.D. (2004). Ternary complex factorserum response factor complex-regulated gene activity is required for cellular proliferation and inhibition of apoptotic cell death. Mol. Cell. Biol. 24, 10340–10351. Wang, J.M., Chao, J.R., Chen, W., Kuo, M.L., Yen, J.J., and Yang-Yen, H.F. (1999). The antiapoptotic gene mcl-1 is up-regulated by the phosphatidylinositol 3-kinase/Akt signaling pathway through a transcription factor complex containing CREB. Mol. Cell. Biol. 19, 6195–6206.

Yamasaki, E., Wada, A., Kumatori, A., Nakagawa, I., Funao, J., Nakayama, M., Hisatsune, J., Kimura, M., Moss, J., and Hirayama, T. (2006). Helicobacter pylori vacuolating cytotoxin induces activation of the proapoptotic protein Bax and Bak, leading to cytochrome c release and cell death, independent of vacuolation. J. Biol. Chem. 281, 11250–11259. Yanai, A., Hirata, Y., Mitsuno, Y., Maeda, S., Shibata, W., Akanuma, M., Yoshida, H., Kawabe, T., and Omata, M. (2003). Helicobacter pylori induces antiapoptosis through Nuclear factor-KB activation. J. Infect. Dis. 188, 1741–1751. Yokoyama, K., Higashi, H., Ishikawa, S., Fujii, Y., Kondo, S., Kato, H., Azuma, T., Wada, A., Hirayama, T., Aburatani, H., and Hatakeyama, M. (2005). Functional antagonism between Helicobacter pylori CagA and vacuolating toxin VacA in control of the NFAT signaling pathway in gastric epithelial cells. Proc. Natl. Acad. Sci. USA 102, 9661– 9666. Zhang, B., Gojo, I., and Fenton, R.G. (2002). Myeloid cell factor-1 is a critical survival factor for multiple myeloma. Blood 99, 1885–1893. Zhu, Y., Wang, C., Huang, J., Ge, Z., Dong, Q., Zhong, X., Su, Y., and Zheng, S. (2007). The Helicobacter pylori virulence factor CagA promotes Erk1/2-mediated Bad phosphorylation in lymphocytes: A mechanism of CagA-inhibited lymphocyte apoptosis. Cell. Microbiol. 9, 952–961.
