INFECTION AND IMMUNITY, Apr. 2003, p. 2153–2162 0019-9567/03/$08.00⫹0 DOI: 10.1128/IAI.71.4.2153–2162.2003 Copyright © 2003, American Society for Microbiology. All Rights Reserved.
Vol. 71, No. 4
Involvement of Myeloid Dendritic Cells in the Development of Gastric Secondary Lymphoid Follicles in Helicobacter pylori-Infected Neonatally Thymectomized BALB/c Mice Toshiki Nishi,1 Kazuichi Okazaki,1* Kimio Kawasaki,1 Toshiro Fukui,1 Hiroyuki Tamaki,1 Minoru Matsuura,1 Masanori Asada,1 Tomohiro Watanabe,1 Kazushige Uchida,1 Norihiko Watanabe,1 Hiroshi Nakase,1 Masaya Ohana,1 Hiroshi Hiai,2 and Tsutomu Chiba1 Department of Gastroenterology & Endoscopic Medicine1 and Department of Pathology,2 Kyoto University, Sakyo, Kyoto, 606-8507, Japan Received 30 September 2002/Returned for modification 25 November 2002/Accepted 27 December 2002
We previously described an animal model of Helicobacter pylori-induced follicular gastritis in neonatally thymectomized (nTx) mice. However, it is still not clear whether antigen-presenting dendritic cells (DCs) in the stomach have a role in the development of secondary follicles in H. pylori-infected nTx mice. We investigated the distribution of DC subsets using this model and examined their roles. To identify lymphoid and myeloid DCs, sections were stained with anti-CD11c (pan-DC marker) in combination with anti-CD8␣ (lymphoid DC marker) or anti-CD11b (myeloid DC marker) and were examined with a confocal microscope. Expression of macrophage inflammatory protein 3␣ (MIP-3␣), which chemoattracts immature DCs, was analyzed by realtime PCR and immunohistochemistry. Follicular dendritic cells (FDCs) were stained with anti-SKY28 antibodies. In noninfected nTx mice, a few myeloid and lymphoid DCs were observed in the bottom portion of the lamina propria, whereas in H. pylori-infected nTx mice, there was an increased influx of myeloid DCs throughout the lamina propria. FDC staining was also observed in the stomachs of members of the infected group. MIP-3␣ gene expression was upregulated in the infected nTx group, and the immunohistochemistry analysis revealed MIP-3␣-positive epithelial cells. These data suggest that H. pylori infection upregulates MIP-3␣ gene expression in gastric epithelial cells and induces an influx of myeloid DCs in the lamina propria of the gastric mucosa in nTx mice. Myeloid DCs and FDCs might contribute to the development of gastric secondary lymphoid follicles in H. pylori-infected nTx mice. Chronic Helicobacter pylori infection induces two distinct types of gastritis, chronic atrophic gastritis and nonatrophic gastritis (4). Nonatrophic gastritis is thought to be a predisposing factor for the development of mucosa-associated lymphoid tissue (MALT) lymphoma (8, 50). The organized structure of the secondary lymphoid tissues is believed to support proper regulation of activation and maturation of the antigen-responsive lymphoid cells (23, 24). Although there is a substantial amount of data on the cellular elements that comprise the lymphoid and nonlymphoid components of the secondary lymphatic organs, little is known about the kinetics and roles of dendritic cells (DCs) that establish and maintain the proper organization of the secondary lymphoid tissues. DCs are immune regulatory cells that not only secrete chemokines and cytokines but also present antigens for T cells (39, 42, 43). In the intestine, DCs are constitutively present in the Peyer’s patches, lamina propria, and mesenteric lymph nodes and have an essential role in the uptake of luminar bacterial antigens (37). Because normal gastric mucosa has no mucosaassociated lymphoid system, very little is known about the role
of DCs in the mucosal immune system of the stomach, especially their role in H. pylori-induced chronic follicular gastritis. In previous studies (33, 46), it was demonstrated that H. pylori infection of BALB/c mice that were thymectomized 3 days after birth (nTx mice) induced follicular gastritis, which resembled human MALT lymphoma. In this model, cooperative involvement of the Th2-type immune response and the Th1 immune response in the development of gastric secondary follicles induced by H. pylori infection was observed. In the present study, to elucidate the involvement of DCs in the development of the follicular formation induced by H. pylori infection, we investigated the presence and characteristics of DCs in the gastric mucosa using the nTx mouse model. In addition, we examined the expression of various chemokines that might influence DCs in the gastric mucosa of nTx mice. MATERIALS AND METHODS Mice and thymectomy. Male and female BALB/c CrSlc mice (Japan SLC, Shizuoka, Japan) were bred in the animal facility at Kyoto University under specific-pathogen-free conditions. Neonatal thymectomies, which induce autoimmune gastritis (AIG) in BALB/c Crslc mice, were performed on day 3 after birth under ether anesthesia, as described previously (33). The results of preliminary studies supported the previously reported findings that there were no immunologic differences between female and male mice (16, 17, 32, 33, 38); we also confirmed that the severity of AIG and the immunologic findings, including the presence of antiparietal autoantibodies and infiltrating lymphocytes, were not different in female and male mice (data not shown). The mice were divided into the following four groups, each of which contained 10 animals: (i) normal
* Corresponding author. Mailing address: Department of Gastroenterology & Endoscopic Medicine, Faculty of Medicine, Kyoto University, 54 Shogoin-Kawara-cho, Sakyo, Kyoto, 606-8507, Japan. Phone: 81-75-751-4319. Fax: 81-75-751-3414. E-mail:
[email protected] .ac.jp. 2153
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INFECT. IMMUN. TABLE 1. Real-time PCR primers and primers and probes
Cytokine or chemokine
INF-␥ IL-4 MIP-3␣ TARC Eotaxin BLC
a
Primer or probe
Sequences (5⬘-3⬘)a
Forward primer Reverse primer Probe Forward primer Reverse primer Probe Forward primer Reverse primer Probe Forward primer Reverse primer Probe Forward primer Reverse primer Probe Forward primer Reverse primer Probe
CTCTGAGACAATGAACGCTACACACT TGGCAGTAACAGCCAGAAACAG FAM-CATCTTGGCTTTGCAGCTTTGCAGCTCTTCCTCATG-TAMRA GGAGCTGCAGAGACTCTTTCG GGCTTTCCAGGAAGTCTTTCAG FAM-CTGCACCATGAATGAGTC CAAGTCCACA-TAMRA CAGAAGCAGCAAGCAACTACGA CTGTCTTGTGAAACCCACAATAGC FAM-TGTTGCCTCTCGTACATACAGACGC-TAMRA GCTGCTGTCCATGGTTTCAAC TTTGTGTTCGCCTGTAGTGCAT FAM-CCACAGAGCAGAAGTCCCTGTTCCCTTT-TAMRA TCACCCTGACTGACCTGTAACTCA CACTTAAAGGCAGAGGCAGGTAA FAM-TGTAGACCAGGCTGACCTCAAACTCACAGA-TAMRA TCCTCGTGCCAAATGGTTACA TAGTGGCTTCAGGCAGCTCTTC FAM-TCTGTCTTCAACTCCCCAAGCTCCAGTG-TAMRA
FAM, 6-carboxyfluorescein; TAMRA, 6-carboxytetramethylrhodamine.
(non-nTx) mice without H. pylori infection, (ii) normal (non-nTx) mice with H. pylori infection, (iii) nTx mice without H. pylori infection, and (iv) nTx mice with H. pylori infection. Bacterial infection of nTx mice. H. pylori TN2FG4, isolated from a Japanese patient with a duodenal ulcer, was donated by M. Nakao (Pharmaceutical Research Division, Takeda Chemical Industries, Ltd., Osaka, Japan). It was maintained in blood agar base no. 2 with horse serum (5%, vol/vol) containing 2.5 mg of amphotericin B per liter, 5 mg of trimethoprim per liter, 1,250 IU of polymyxin B per liter, and 10 mg of vancomycin per liter. The plates were incubated in a microaerophilic atmosphere at 37°C for 48 h. The inoculated H. pylori strain, TN2FG4, was CagA and VacA positive, as described elsewhere (49). Both nTx and non-nTx mice that were 6 weeks old were orally infected with 108 H. pylori cells as described previously (33). Infected mice were sacrificed 1, 4, 8, and 12 weeks later. Noninfected mice were sacrificed at the same time. Colonization by H. pylori was confirmed by Giemsa staining, culture analysis, and PCR Southern blot analysis for the urease gene by using DNA extracted from the corpus mucosa. The urease gene was amplified 12 weeks after infection for all of the infected mice (33). After decapitation, the sera and stomachs were immediately frozen and stored until they were used. The abnormal hyperimmune status of the nTx mice was confirmed by measuring the serum levels of autoantibodies against parietal cells by an enzyme-linked immunosorbent assay immediately before infection (6 weeks after the mice were thymectomized), as described previously (33). Histology. The stomach was removed from each mouse, fixed with 4% phosphate-buffered formaldehyde (pH 7.2), and prepared for histologic examination. The sections were stained with hematoxylin and eosin. Double immunofluorescence staining for DC. The immunohistochemistry analysis was performed with serially cut frozen sections by using antibodies (Abs) reactive to DCs (for the pan-DC marker, fluorescein isothiocyanate–anti-CD11c; for the linear DC marker, biotin–anti-CD11b and -CD8␣; Pharmingen, San Diego, Calif.), to major histocompatibility complex class II (biotin–anti-Iab; Pharmingen), to costimulatory molecules (biotin–anti-CD80 and -CD86; Pharmingen), and to macrophage inflammatory protein 3␣ (MIP-3␣) (biotin–anti-MIP3␣; R&D Systems, Minneapolis, Minn.). Briefly, freshly frozen sections were fixed in acetone for 10 min, rinsed in phosphate-buffered saline (PBS) (pH 7.2), and incubated with 10% normal rat, hamster, or mouse serum (as a blocking agent) for 20 min. Sections were incubated for 1 h with biotin-labeled Abs. After incubation, the sections were washed with PBS and then incubated with Texas Red avidin (Vector Laboratories, Inc., Burlingame, Calif.) for 30 min. The sections were washed with PBS and incubated with 10% normal hamster serum (as a blocking agent) for 20 min and incubated for 1 h with the fluorescein isothiocyanate-labeled monoclonal Abs. After incubation, the sections were washed with PBS and then washed with distilled water and mounted on glass slides with Vectashield (Vector Laboratories, Inc.). Sections were viewed with a Zeiss Axioplan/Bio-Rad MRC 1024 confocal laser microscope with a 10⫻ objective. Control sections were exposed to
normal rat serum instead of monoclonal Abs. There was no staining in the control sections. Immunohistochemistry analysis for follicular dendritic cells (FDCs). Frozen sections prepared from each stomach were stained by the avidin-biotin complex method. Anti-mouse SKY28 (12) Ab, kindly provided by N. Imazeki (National Defense Medical College, Tokorozawa, Japan), or isotype-matched control goat immunoglobulin G was used as the primary Ab. Rapid immunohistochemical staining of the primary Ab was performed by using a Vectastain universal quick kit (Vector Laboratories, Inc.) according to the manufacturer’s instructions. Localization of the antigens was visualized by incubation with a diaminobenzidine solution, and the antigens were counterstained with methyl green. Measurement of the number of DC subpopulations. Myeloid DCs and lymphoid DCs were quantified in four different areas of the gastric mucosa (surface epithelium, upper portion of the lamina propria, middle portion of the lamina propria, bottom portion of the lamina propria) of both 14-week-old nTx mice that were not infected with H. pylori and 14-week-old nTx mice that were infected (8 weeks postinfection). The total number of positive cells in five continuous gastric gland units was determined for five different regions in one frozen section. Each group consisted of five animals, and the mean ⫾ standard error of the mean was calculated for each group. Chemokine and cytokine expression. Total RNA was extracted from the stomachs of 10 mice in each group by using the single-step guanidium thiocyanatephenol-chloroform method. The extracted RNA preparations were reverse transcribed with MultiScribe reverse transcriptase (PE Applied Biosystems, Foster City, Calif.). The resultant cDNAs (50 ng/reaction mixture) were analyzed for expression of gamma interferon (IFN-␥), interleukin-4 (IL-4), MIP-3␣, B-lymphocyte chemoattractant (BLC), thymus- and activation-regulated chemokine (TARC), and eotaxin genes by using the TaqMan PCR assay with an ABI Prism 7700 sequence detection system (Perkin-Elmer, Foster City, Calif.). The reaction mixtures were incubated for 2 min at 50°C, denatured for 10 min at 95°C, and subjected to 40 two-step amplification cycles consisting of annealing and extension at 60°C for 1 min followed by denaturation at 95°C for 15 s. The primers and probes for cytokines and chemokines designed with the PRIMER-EXPRESS software (PE Applied Biosystems) are shown in Table 1. Reaction mixtures were incubated for 2 min at 50°C, denatured for 10 min at 95°C, and subjected to 45 amplification cycles consisting of annealing and extension at 60°C for 1 min followed by denaturation at 95°C for 15 s. All TaqMan PCR data were captured by using the Sequence Detector software (PE Applied Biosystems). The template concentration in each reaction mixture was determined by comparison with a gene-specific standard curve constructed by using the cDNA of positive control samples and normalized by dividing the number of copies of the target gene per nanogram by the number of copies of a housekeeping gene (glyceraldehyde-3-phosphate dehydrogenase [GAPDH] gene, amplified by using TaqMan rodent GAPDH control reagents; PE Applied Biosystems) per nanogram and was expressed as a percentage (mean ⫾ standard error).
DCs IN H. PYLORI-INDUCED FOLLICULAR GASTRITIS
VOL. 71, 2003 Statistical analysis. All results are expressed as the mean ⫾ standard error for each sample. The differences in the quantitative mRNA expression of cytokines and chemokines were analyzed by using Wilcoxon’s t test. A two-tailed P value of less than 0.05 was considered to be statistically significant.
RESULTS Double immunofluorescence staining for DCs. There was no apparent gastritis in the stomachs of normal BALB/c mice (non-nTx, 6 weeks old) (Fig. 1a), and there was no staining for either myeloid or lymphoid DCs throughout the experiment (Fig. 1b and c). H. pylori-infected non-nTx mice were similar to noninfected non-nTx mice; i.e., there was no apparent gastritis and no DC staining (data not shown). For 6-week-old noninfected nTx mice, the histologic findings obtained for the stomachs indicated that there was mild gastritis (Fig. 1d), and there were a few stained myeloid DCs (Fig. 1e) but no stained lymphoid DCs (Fig. 1f). For 14- and 18week-old noninfected nTx mice, the histologic findings obtained for the stomachs were typical of AIG, showing a loss of parietal cells, proliferation of pit cells, and moderate to severe lymphocyte infiltration (Fig. 1g and j). There were a few lymphoid and myeloid DCs in the bottom portion of the lamina propria in the stomachs of both 14- and 18-week-old noninfected nTx mice (Fig. 1h, i, k, and l). For H. pylori-infected nTx mice, the histologic findings were similar to those obtained for noninfected nTx mice for the first 12 weeks after infection (Fig. 2a, d, and g). At 12 weeks after infection, however, follicle formation was observed (Fig. 2j). On the other hand, lymphoid DCs were observed in the bottom portion of the lamina propria and submucosal area (Fig. 2c, f, i, and l) after 1 week of infection; this pattern continued during the observation period, but it was not different from that in noninfected nTx mice (Table 2). In contrast, H. pylori infection remarkably enhanced the influx of myeloid DCs in nTx mice (Fig. 2 and Table 2). At 1 week after infection, myeloid DCs were observed throughout the lamina propria, and the number of these cells increased thereafter (Fig. 2b, e, and h). Twelve weeks after infection, myeloid DCs were induced around the follicles (Fig. 2k). Eight weeks after infection, the number of myeloid DCs in the infected nTx mice was significantly greater than that in noninfected nTx mice, and moreover, the number of myeloid DCs in H. pylori-infected nTx mice was also significantly higher than the number of lymphoid DCs in the lamina propria of the same mice except for the bottom portion (Table 2). In the stomachs of non-H. pylori-infected nTx mice, there was not a significant difference between the number of myeloid DCs and the number of lymphoid DCs (Table 2). There was no correlation between the number of myeloid DCs and H. pylori colonization (data not shown). Detection of activated DCs by immunohistochemistry analysis. To investigate activation of DCs in the gastric mucosa of H. pylori-infected nTx mice (8 weeks postinfection, 14 weeks old) and noninfected nTx mice (14 weeks old), we performed double immunofluorescence staining using mouse anti-CD11c for a pan-DC marker and mouse anti-IAb, -CD80, and -CD86 for activated markers. There were a few activated DCs in the bottom portion of the lamina propria of the H. pylori-infected stomachs (Fig. 3d, e, and f). In contrast, there was no staining
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of activated DCs in the stomachs of the noninfected nTx mice (Fig. 3a, b, and c). Analysis of Th1/Th2 balance. The cytokine and chemokine mRNA profiles for the gastric mucosa of noninfected normal mice (non-nTx, 14 weeks old), noninfected nTx mice (14 weeks old), and H. pylori-infected nTx mice (8 weeks after infection, 14 weeks old) were determined by real-time PCR. Although the nTx mice had significantly enhanced IFN-␥ gene expression (0.3 ⫾ 0.1 versus 3.1 ⫾ 0.9 copies/GAPDH gene copy; P ⬍ 0.05) compared with non-nTx normal mice, there was no significant difference between the levels of IFN-␥ gene expression in the H. pylori-infected and noninfected nTx mouse groups (3.2 ⫾ 0.8 versus 3.1 ⫾ 0.9 copies/GAPDH gene copy) (Fig. 4a). On the other hand, IL-4 gene expression was not affected by the nTx procedure in noninfected mice (0.03 ⫾ 0.01 versus 0.1 ⫾ 0.1 copies/GAPDH gene copy). IL-4 gene expression, however, was significantly enhanced by H. pylori infection in the nTx mice (0.8 ⫾ 0.1 versus 0.1 ⫾ 0.1 copies/GAPDH gene copy; P ⬍ 0.01) (Fig. 4b). The levels of expression of the genes encoding two representative Th2 chemokines, TARC and eotaxin, were also significantly increased by H. pylori infection in nTx mice (4.5 ⫾ 0.9 versus 1.5 ⫾ 0.9 copies/GAPDH gene copy [P ⬍ 0.05] and 116.0 ⫾ 5.6 versus 36.6 ⫾ 26.1 copies/GAPDH gene copy [P ⬍ 0.05], respectively) (Fig. 4c and d), although nTx alone did not affect the expression of the genes encoding these chemokines. Analysis of MIP-3␣ expression. We analyzed expression of the gene encoding MIP-3␣, which chemoattracts immature DCs (2, 22), in the gastric mucosa of non-H. pylori-infected non-nTx mice (control, 14 weeks old), non-H. pylori-infected nTx mice (14 weeks old), and H. pylori-infected nTx mice (8 weeks postinfection, 14 weeks old). Although nTx alone did not have an effect, H. pylori infection significantly enhanced MIP-3␣ mRNA expression in nTx mice (4.0 ⫾ 1.4 versus 0.06 ⫾ 0.03 copies/GAPDH gene copy; P ⬍ 0.05) (Fig. 4e). In the gastric mucosa of control mice, there was no staining of DCs or MIP-3␣ 14 weeks after birth (Fig. 5a). MIP-3␣ staining, however, was observed after H. pylori infection in the gastric mucosa of nTx mice, as well as non-nTx mice (Fig. 5b and d). On the other hand, in the stomachs of non-H. pyloriinfected nTx mice, only DCs were stained (Fig. 5c). Immunohistochemistry analysis of FDCs and BLC gene expression. To detect FDCs in the gastric mucosa, we used monoclonal Ab SKY28 against mouse FDCs, which was obtained from FDC cluster-enriched cells as an immunogen (12). There were anti-mouse SKY28 Ab-positive cells in the gastric mucosa of H. pylori-infected nTx mice (8 weeks postinfection, 14 weeks old) (Fig. 6c), whereas there were no positive cells in the stomachs of noninfected nTx mice (14 weeks old) (Fig. 6b). In H. pylori-infected nTx mice, expression of the gene encoding BLC, a chemokine produced by FDCs (9, 28) that chemoattracts B cells (9, 21), was significantly elevated compared to expression in noninfected nTx mice (0.014 ⫾ 0.006 versus 0.001 ⫾ 0.001 copies/GAPDH gene copy; P ⬍ 0.05) (Fig. 4f). DISCUSSION It was previously reported that H. pylori infection induces secondary lymphoid follicle formation in the gastric mucosa of nTx BALB/c mice (33, 46). Various factors are involved in the
FIG. 1. Confocal microscopic analysis of myeloid and lymphoid DCs in the stomach and hematoxylin and eosin staining of non-nTx (normal) and nTx mice. Two-color staining was performed with 6-m frozen stomach sections by using Abs against CD11c (green) in combination with anti-CD11b (red) (b, e, h, and k) or anti-CD8␣ (red) (c, f, i, and l). Sections were prepared from 6-week-old noninfected normal mice (a to c), 6-week-old noninfected nTx mice (d to f), 14-week-old noninfected nTx mice (g to i), and 18-week-old noninfected nTx mice (j to l). DCs positive for either CD11b or CD8␣ were stained yellow. Sections of the same animals were stained with hematoxylin and eosin (a, d, g, and j) to visualize stomach morphology, and the presence of gastritis in nTx mice was confirmed. There was no apparent gastritis (a) and no DC staining (b and c) in normal mice. The histologic findings for the stomachs of nTx mice were typical of AIG, with a loss of parietal cells, proliferation of pit cells, and moderate to severe lymphocyte infiltration (d, g, and j). In 6-week-old nTx mice, staining of a few myeloid DCs was observed at the bottom of the lamina propria (e), whereas there was no lymphoid DC staining (f). In 14- and 18-week-old nTx mice, some staining of both myeloid and lymphoid DCs was observed in the bottom portion of the lamina propria. 2156
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FIG. 2. Confocal microscopic analysis of myeloid and lymphoid DCs in the stomach and hematoxylin and eosin staining of H. pylori-infected nTx mice. The histologic findings for the stomachs from H. pylori-infected nTx mice were mild to severe gastritis (a, d, and g [1, 4, and 8 weeks after infection, respectively]). At 12 weeks after infection (18-week-old mice), there was follicle formation (j). Staining of lymphoid DCs (CD11c and CD8␣ double positive; yellow) was observed in the bottom portion of the lamina propria and submucosal area throughout the experiment (c, f, i, and l). In contrast, there were myeloid DCs (CD11c and CD11b double positive; yellow) throughout the lamina propria, and the number increased linearly during the observation period (b, e, and h). In H. pylori-infected nTx mice at 12 weeks after infection (18 weeks old), myeloid DCs were induced around the follicle (k).
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TABLE 2. Numbers of DCs in the gastric mucosa of nTx mice that were infected or not infected with H. pylori No. ina: DCs
Myeloid
Lymphoid
Location
Noninfected nTx mice
H. pylori-infected nTx mice
Surface epithelium Upper portion of lamina propria Middle portion of lamina propria Bottom portion of lamina propria
0.2 ⫾ 0.1 0.1 ⫾ 0.1
0.7 ⫾ 0.2b,c 5.0 ⫾ 0.7b,c
0.3 ⫾ 0.2
1.9 ⫾ 0.4b,c
2.9 ⫾ 0.4
2.4 ⫾ 0.3
Surface epithelium Upper portion of lamina propria Middle portion of lamina propria Bottom portion of lamina propria
0.1 ⫾ 0.1 0.0
0.1 ⫾ 0.1 0.0
0.1 ⫾ 0.1
0.2 ⫾ 0.1
2.0 ⫾ 0.2
1.8 ⫾ 0.3
a The total number of positive cells in five continuous gastric gland units was counted in five different regions in one frozen section at 14 weeks after birth (8 weeks after H. pylori infection). Each group consisted of five animals, and the mean ⫾ standard error of the mean was calculated for each group. b Significantly greater than corresponding values for noninfected nTx mice (P ⬍ 0.01). c Significantly greater than the number of lymphoid DCs in the same location (P ⬍ 0.01).
organogenesis of lymphoid tissues in the developmental stage (1, 11, 18, 19, 27, 34), but little is known about the formation of lymphoid follicles in H. pylori-infected stomachs. Although there are similarities and differences between H. pylori infection in the human stomach and H. pylori infection in the murine stomach, the bacterial strain and host factors have critical roles in the development of follicular gastritis. In humans, the
prevalence of follicular gastritis in H. pylori-infected patients has been reported to vary from 24 to 72% (3, 5, 7, 20, 44, 50–52), probably due to differences in H. pylori strains and ethnic groups. In murine models, H. pylori-induced gastritis depends both on the bacterial strain and the host (48). Previous studies showed that CagA-positive H. pylori infection induced follicular gastritis in nTx BALB/c mice but not in nonnTx normal BALB/c mice or nTx mice belonging to other strains (33, 46). In terms of immunological factors, the roles of DCs in the development of follicles in H. pylori-infected stomachs are still unclear. For humans, only one report has described the presence of a DC subpopulation in an H. pyloripositive MALT lymphoma (40). In the present study, there were no DCs in the gastric mucosa of normal mice, as has been suggested by other workers (31), and only a few myeloid and lymphoid DCs were present in the bottom portion of the lamina propria and in the submucosal area of the gastric mucosa of nTx mice that were not infected with H. pylori. The most noteworthy finding in this study was that in contrast to the noninfected nTx mice, a large number of myeloid DCs were diffusely infiltrated throughout the lamina propria in the H. pylori-infected nTx mouse stomachs, although the degree of lymphoid DC infiltration was not changed by H. pylori infection. These data indicate that myeloid DCs might have an important role in the development of gastric secondary lymphoid follicles induced by H. pylori infection in nTx mice. Different DC subsets, such as lymphoid and myeloid DCs, induce distinct immune responses (Th1 and Th2) (30). In mice, freshly isolated CD8␣⫹ (lymphoid) and CD8␣⫺ (myeloid) DCs from spleens (25, 36) or Peyer’s patches (13) induce Th1 and Th2 responses, respectively. CD8␣⫹ DCs secrete IL-12, an essential cytokine for inducing the Th1-type immune response, whereas CD8␣⫺ DCs secrete IL-4 and IL-10, which elicit Th2
FIG. 3. Confocal microscopic analysis of activated DCs in the stomachs of H. pylori-infected and noninfected nTx mice (12 weeks after infection, 18 weeks old). Two-color staining was performed with sections by using Abs against CD11c (green) in combination with anti-IAb (red) (a and d), anti-CD80 (red) (b and e), or anti-CD86 (red) (c and f). In the stomachs of noninfected nTx mice, there was no staining of double-positive cells (a, b, and c). In contrast, there were a few double-positive cells in the bottom portions of the lamina propria in the stomachs of H. pylori-infected nTx mice (yellow) (d, e, and f) (the small boxes were magnified to produce the large boxes).
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FIG. 4. Profiles of cytokine and chemokine mRNA expression in the gastric mucosa of non-H. pylori-infected normal mice (non-nTx, 14 weeks old), non-H. pylori-infected nTx mice (14 weeks old), and H. pylori-infected nTx mice (8 weeks after infection, 14 weeks old), as determined by real-time PCR. Ten mice from each group were analyzed. The data are expressed as the means ⫾ standard errors for the number of copies of the mRNA per GAPDH gene copy. One asterisk and two asterisks indicate that the data were significantly different at P values of ⬍0.05 and ⬍0.01, respectively. NS, not significant; Hp, H. pylori infected.
immunity (14, 25, 35, 41). Neonatal thymectomy induces AIG in BALB/c mice, and Th1 cells predominantly infiltrate the gastric mucosa, while Th2 cells reside mainly in the regional lymph nodes and not in the lamina propria in nTx mice (17). In contrast, it was demonstrated previously that secondary lymph follicle formation induced by H. pylori infection in nTx mice was associated with activation of both the Th1 and Th2 immune responses (46). Moreover, we demonstrated in the
present study that in addition to the level of IL-4 gene expression, the levels of expression of the genes encoding the CC chemokines eotaxin and TARC, both of which evoke chemotaxis of the Th2-type CD4⫹ T lymphocytes (22, 39), were significantly increased in the stomachs of H. pylori-infected nTx mice. These data suggest that myeloid DCs have an important role in the induction of the Th2 response by H. pylori infection in nTx mouse stomachs.
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FIG. 5. Confocal microscopic analysis of DCs and MIP-3␣ staining in the gastric mucosa of non-H. pylori-infected normal mice (non-nTx) (a), H. pylori-infected normal mice (non-nTx, 14 weeks old) (b), non-H. pylori-infected nTx mice (14 weeks old) (c), and H. pylori-infected nTx mice (8 weeks after infection, 14 weeks old) (d). Two-color staining of the sections was performed with Ab against CD11c (green) in combination with anti-MIP-3␣ (red).
On the other hand, Helicobacter infection upregulates IFN-␥ expression but not IL-4 expression in the stomachs of C57BL/6 mice (29). The differences between mouse strains might be due to differences in the genetic backgrounds of the Th1-dominated C57BL/6 mice and the Th2-dominated BALB/c mice (10). MIP-3␣, which chemoattracts CCR6-expressing immature DCs (2, 22), is abundantly expressed in mouse and human inflammatory enteric mucosa (15, 45). Iwasaki and Kelsall (13) recently demonstrated that in Peyer’s patches a subset of CD11b⫹ myeloid DCs that is positive for CCR6 lines the epithelial layer of the dome and is the only DC population that responds to MIP-3␣. Therefore, we examined whether MIP-3␣ is involved in the preferential increase in the number of myeloid DCs in the gastric mucosa in our mouse model, and our immunohistochemistry analysis revealed that MIP-3␣ was present in the epithelial cells of the stomachs of H. pyloriinfected nTx mice, whereas there was no MIP-3␣ in non-H. pylori-infected nTx or non-nTx mice. These findings indicate that H. pylori infection is responsible for MIP-3␣ induction in the epithelial cells and that MIP-3␣ expressed in gastric epithelial cells might have an important role in the migration of myeloid DCs into the upper portion of the lamina propria. MIP-3␣ expression was observed not only in nTx mice but also in non-nTx mice that were infected with H. pylori, in which we did not observe significant infiltration of myeloid DCs into the lamina propria or secondary lymph follicles. These results sug-
gest that MIP-3 ␣ is necessary, but not sufficient, for the preferential infiltration of myeloid DCs into the lamina propria with resulting follicular formation induced by H. pylori infection and that, in addition to MIP-3␣, other factors are required for such responses to H. pylori infection in nTx mice. In human tonsils, MCP-4 molecules expressed on basal epithelial cells lining the blood vessels have an important role in mobilizing circulatory DCs into the inflammatory site of a tonsil (47). Although similar molecules have not been identified yet in mice, H. pylori infection might induce an unknown chemokine similar to MCP-4, which chemoattracts myeloid DCs from the bloodstream to the inflamed gastric mucosa. In the present study, activated DCs were present in the lamina propria of H. pylori-infected nTx mice, whereas there were no such cells in noninfected nTx mice. These findings suggest that DCs were activated by H. pylori infection. It should be noted that the activated DCs were localized in the bottom portion of the lamina propria. Thus, the activated DCs might migrate to the regional lymph nodes. Another interesting observation is that localization of the activated DCs were superimposed on lymphoid DCs. Recently, Martinez del Hoyo et al. (26) reported that lymphoid and myeloid DCs represent different maturation or differentiation stages of the same DC population and that lymphoid DCs represent the late stage of DC differentiation. Therefore, although the number of lymphoid DCs was not increased by H. pylori infection, an interesting question is whether the lymphoid DCs distributed over
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nificantly greater in the stomachs of H. pylori-infected nTx mice than in the stomachs of noninfected nTx mice. In addition, we confirmed that FDCs, which secrete BLC (9, 28), appeared in the stomachs of H. pylori-infected nTx mice but not in the stomachs of noninfected nTx mice. These findings suggest that BLC has a critical role in maintaining secondary lymphoid follicles. In summary, the present study demonstrated that H. pylori infection induces MIP-3␣ expression in gastric epithelial cells and promotes preferential migration of myeloid DCs into the lamina propria in nTx mice. This effect, coordinated with FDCs, might promote Th2 immune responses, which might then facilitate the development of secondary lymphoid follicles in H. pylori-infected gastric mucosa. ACKNOWLEDGMENTS This study was supported by grant-in-aid for scientific research (C) 14570463 from the Ministry of Culture and Science of Japan, by grantin-aid JSPS-RFTF97I00201 for the Research for the Future Program from the Japan Society for the Promotion of Science, by research funds from the Japanese Foundation for Research and Promotion of Endoscopy (grant JFE-2001), and by the Shimidzu Immunology Foundation, 2000. REFERENCES
FIG. 6. Immunohistochemical staining of FDCs in the gastric mucosa of nTx mice (original magnification, ⫻400). Gastric tissues were incubated with isotype-matched control goat immunoglobulin G (a) and anti-mouse SKY28 Ab (b and c). Positive cells (arrows) were observed in the stomachs of H. pylori-infected nTx mice (8 weeks after infection, 14 weeks old) (c), whereas there were no positive cells in the stomachs of noninfected nTx mice (14 weeks old) (b).
the bottom portion of the lamina propria in H. pylori-infected nTx mice are mature DCs. In previous studies (33, 46), increases in the number of B cells were observed along with the formation of germinal centers induced by H. pylori infection in nTx mice. One candidate factor that could attract B cells to follicles is the potent chemoattractant BLC, which is constitutively expressed in lymphoid tissues (9, 21). Mice lacking CXCR5, the BLC receptor, exhibit a defect in the development of B-cell follicles in the spleen, Peyer’s patches, and inguinal lymph nodes (6). Thus, a BLC-CXCR5 interaction might have an important role in homing B cells into secondary lymphoid organs. The present study demonstrated that expression of the BLC gene was sig-
1. De Togni, P., J. Goellner, N. H. Ruddle, P. R. Streeter, A. Fick, S. Mariathasan, S. C. Smith, R. Carlson, L. P. Shornick, J. Strauss-Schoenberger, et al. 1994. Abnormal development of peripheral lymphoid organs in mice deficient in lymphotoxin. Science 264:703–707. 2. Dieu-Nosjean, M. C., A. Vicari, S. Lebecque, and C. Caux. 1999. Regulation of dendritic cell trafficking: a process that involves the participation of selective chemokines. J. Leukoc. Biol. 66:252–262. 3. Di Napoli, A., R. Petrino, M. Boero, D. Bellis, and L. Chiandussi. 1992. Quantitative assessment of histological changes in chronic gastritis after eradication of Helicobacter pylori. J. Clin. Pathol. 45:796–798. 4. Dixon, M. F., R. M. Genta, J. H. Yardley, and P. Correa. 1996. Classification and grading of gastritis. The Updated Sydney Systematic International Workshop on the Histopathology of Gastritis, Houston 1994. Am. J. Surg. Pathol. 20:1161–1181. 5. Eidt, S., and M. Stolte. 1993. Prevalence of lymphoid follicles and aggregates in Helicobacter pylori gastritis in antral and body mucosa. J. Clin. Pathol. 46:832–835. 6. Forster, R., A. E. Mattis, E. Kremmer, E. Wolf, G. Brem, and M. Lipp. 1996. A putative chemokine receptor, BLR1, directs B cell migration to defined lymphoid organs and specific anatomic compartments of the spleen. Cell. 87:1037–1047. 7. Genta, R. M., and H. W. Hamner. 1994. The significance of lymphoid follicles in the interpretation of gastric biopsy specimens. Arch. Pathol. Lab. Med. 118:740–743. 8. Genta, R. M., H. W. Hamner, and D. Y. Graham. 1993. Gastric lymphoid follicles in Helicobacter pylori infection: frequency, distribution, and response to triple therapy. Hum. Pathol. 24:577–583. 9. Gunn, M. D., V. N. Ngo, K. M. Ansel, E. H. Ekland, J. G. Cyster, and L. T. Williams. 1998. A B-cell-homing chemokine made in lymphoid follicles activates Burkitt’s lymphoma receptor-1. Nature 391:799–803. 10. Heinzel, F. P., M. D. Sadick, B. J. Holaday, R. L. Coffman, and R. M. Locksley. 1989. Reciprocal expression of interferon gamma or interleukin 4 during the resolution or progression of murine leishmaniasis. Evidence for expansion of distinct helper T cell subsets. J. Exp. Med. 169:59–72. 11. Hikida, M., Y. Nakayama, Y. Yamashita, Y. Kumazawa, S. I. Nishikawa, and H. Ohmori. 1998. Expression of recombination activating genes in germinal center B cells: involvement of interleukin 7 (IL-7) and the IL-7 receptor. J. Exp. Med. 188:365–372. 12. Imazeki, N., A. Takeuchi, A. Senoo, and Y. Fuse. 1994. New monoclonal antibodies directed against mouse follicular dendritic cells. J. Histochem. Cytochem. 42:329–335. 13. Iwasaki, A., and B. L. Kelsall. 2000. Localization of distinct Peyer’s patch dendritic cell subsets and their recruitment by chemokines macrophage inflammatory protein (MIP)-3alpha, MIP-3beta, and secondary lymphoid organ chemokine. J. Exp. Med. 191:1381–1394. 14. Iwasaki, A., and B. L. Kelsall. 2001. Unique functions of CD11b⫹, CD8 alpha⫹, and double-negative Peyer’s patch dendritic cells. J. Immunol. 166: 4884–4890. 15. Izadpanah, A., M. B. Dwinell, L. Eckmann, N. M. Varki, and M. F. Kagnoff.
2162
16.
17. 18.
19.
20.
21.
22.
23. 24. 25.
26.
27. 28.
29.
30. 31.
32.
33.
NISHI ET AL.
2001. Regulated MIP-3alpha/CCL20 production by human intestinal epithelium: mechanism for modulating mucosal immunity. Am. J. Physiol. Gastrointest. Liver Physiol. 280:G710–G719. Jones, C. M., J. M. Callaghan, P. A. Gleeson, Y. Mori, T. Masuda, and B. H. Toh. 1991. The parietal cell autoantigens recognized in neonatal thymectomy-induced murine gastritis are the alpha and beta subunits of the gastric proton pump. Gastroenterology 101:287–294. (Corrected.) Katakai, T., K. J. Mori, T. Masuda, and A. Shimizu. 1998. Differential localization of Th1 and Th2 cells in autoimmune gastritis. Int. Immunol. 10:1325–1334. Koni, P. A., R. Sacca, P. Lawton, J. L. Browning, N. H. Ruddle, and R. A. Flavell. 1997. Distinct roles in lymphoid organogenesis for lymphotoxins alpha and beta revealed in lymphotoxin beta-deficient mice. Immunity 6:491–500. Korner, H., M. Cook, D. S. Riminton, F. A. Lemckert, R. M. Hoek, B. Ledermann, F. Kontgen, B. Fazekas de St Groth, and J. D. Sedgwick. 1997. Distinct roles for lymphotoxin-alpha and tumor necrosis factor in organogenesis and spatial organization of lymphoid tissue. Eur. J. Immunol. 27: 2600–2609. Ladas, S. D., T. Rokkas, S. Georgopoulos, P. Kitsanta, C. Liatsos, P. Eustathiadou, A. Karameris, C. Spiliadi, and S. A. Raptis. 1999. Predictive factors and prevalence of follicular gastritis in adults with peptic ulcer and nonulcer dyspepsia. Dig. Dis. Sci. 44:1156–1160. Legler, D. F., M. Loetscher, R. S. Roos, I. Clark-Lewis, M. Baggiolini, and B. Moser. 1998. B cell-attracting chemokine 1, a human CXC chemokine expressed in lymphoid tissues, selectively attracts B lymphocytes via BLR1/ CXCR5. J. Exp. Med. 187:655–660. Liao, F., R. L. Rabin, C. S. Smith, G. Sharma, T. B. Nutman, and J. M. Farber. 1999. CC-chemokine receptor 6 is expressed on diverse memory subsets of T cells and determines responsiveness to macrophage inflammatory protein 3 alpha. J. Immunol. 162:186–194. Liu, Y. J., G. Grouard, O. de Bouteiller, and J. Banchereau. 1996. Follicular dendritic cells and germinal centers. Int. Rev. Cytol. 166:139–179. MacLennan, I. C. 1994. Germinal centers. Annu. Rev. Immunol. 12:117–139. Maldonado-Lopez, R., T. De Smedt, P. Michel, J. Godfroid, J., Pajak, C. Heirman, K. Thielemans, O. Leo, J. Urbain, and M. Moser. 1999. CD8alpha⫹ and CD8alpha⫺ subclasses of dendritic cells direct the development of distinct T helper cells in vivo. J. Exp. Med. 189:587–592. Martinez del Hoyo, G., P. Martin, C. F. Arias, A. R. Marin, and C. Ardavin. 2002. CD8alpha⫹ dendritic cells originate from the CD8alpha⫺ dendritic cell subset by a maturation process involving CD8alpha, DEC-205, and CD24 up-regulation. Blood 99:999–1004. Matsumoto, M., S. Mariathasan, M. H. Nahm, F. Baranyay, J. J. Peschon, and D. D. Chaplin. 1996. Role of lymphotoxin and the type I TNF receptor in the formation of germinal centers. Science 271:1289–1291. Mazzucchelli, L., A. Blaser, A. Kappeler, P. Scharli, J. A. Laissue, M. Baggiolini, and M. Uguccioni. 1999. BCA-1 is highly expressed in Helicobacter pylori-induced mucosa-associated lymphoid tissue and gastric lymphoma. J. Clin. Investig. 104:R49–R54. Mohammadi, M., S. Czinn, R. Redline, and J. Nedrud. 1996. Helicobacterspecific cell-mediated immune responses display a predominant Th1 phenotype and promote a delayed-type hypersensitivity response in the stomachs of mice. J. Immunol. 156:4729–4738. Moser, M., and K. M. Murphy. 2000. Dendritic cell regulation of TH1-TH2 development. Nat. Immunol. 1:199–205. Ninomiya, T., H. Matsui, S. M. Akbar, H. Murakami, and M. Onji. 2000. Localization and characterization of antigen-presenting dendritic cells in the gastric mucosa of murine and human autoimmune gastritis. Eur. J. Clin. Investig. 30:350–358. Nishio, A., M. Hosono, Y. Watanabe, M. Sakai, M. Okuma, and T. Masuda. 1994. A conserved epitope on H⫹,K(⫹)-adenosine triphosphatase of parietal cells discerned by a murine gastritogenic T-cell clone. Gastroenterology 107:1408–1414. Oshima, C., K. Okazaki, Y. Matsushima, M. Sawada, T. Chiba, K. Takahashi, H. Hiai, T. Katakai, S. Kasakura, and T. Masuda. 2000. Induction of follicular gastritis following postthymectomy autoimmune gastritis in Helicobacter pylori-infected BALB/c mice. Infect. Immun. 68:100–106.
Editor: A. D. O’Brien
INFECT. IMMUN. 34. Pasparakis, M., L. Alexopoulou, M. Grell, K. Pfizenmaier, H. Bluethmann, and G. Kollias. 1997. Peyer’s patch organogenesis is intact yet formation of B lymphocyte follicles is defective in peripheral lymphoid organs of mice deficient for tumor necrosis factor and its 55-kDa receptor. Proc. Natl. Acad. Sci. USA 94:6319–6323. 35. Pulendran, B., J. Lingappa, M. K. Kennedy, J. Smith, M. Teepe, A. Rudensky, C. R. Maliszewski, and E. Maraskovsky. 1997. Developmental pathways of dendritic cells in vivo: distinct function, phenotype, and localization of dendritic cell subsets in FLT3 ligand-treated mice. J. Immunol. 159:2222– 2231. 36. Pulendran, B., J. L. Smith, G. Caspary, K. Brasel, D. Pettit, E. Maraskovsky, and C. R. Maliszewski. 1999. Distinct dendritic cell subsets differentially regulate the class of immune response in vivo. Proc. Natl. Acad. Sci. USA 96:1036–1041. 37. Rescigno, M., M. Urbano, B. Valzasina, M. Francolini, G. Rotta, R. Bonasio, F. Granucci, J. P. Kraehenbuhl, and P. Ricciardi-Castagnoli. 2001. Dendritic cells express tight junction proteins and penetrate gut epithelial monolayers to sample bacteria. Nat. Immunol. 2:361–367. 38. Sakaguchi, S., K. Fukuma, K. Kuribayashi, and T. Masuda. 1985. Organspecific autoimmune diseases induced in mice by elimination of T cell subset. I. Evidence for the active participation of T cells in natural self-tolerance; deficit of a T cell subset as a possible cause of autoimmune disease. J. Exp. Med. 161:72–87. 39. Sallusto, F., D. Lenig, C. R. Mackay, and A. Lanzavecchia. 1998. Flexible programs of chemokine receptor expression on human polarized T helper 1 and 2 lymphocytes. J. Exp. Med. 187:875–883. 40. Sarsfield, P., D. B. Jones, A. C. Wotherspoon, T. Harvard, and D. H. Wright. 1996. A study of accessory cells in the acquired lymphoid tissue of helicobacter gastritis. J. Pathol. 180:18–25. 41. Sousa, C. R., S. Hieny, T. Scharton-Kersten, D. Jankovic, H. Charest, R. N. Germain, and A. Sher. 1997. In vivo microbial stimulation induces rapid CD40 ligand-independent production of interleukin 12 by dendritic cells and their redistribution to T cell areas. J. Exp. Med. 186:1819–1829. 42. Steinman, R. M., M. Pack, and K. Inaba. 1997. Dendritic cell development and maturation. Adv. Exp. Med. Biol. 417:1–6. 43. Steinman, R. M., M. Pack, and K. Inaba. 1997. Dendritic cells in the T-cell areas of lymphoid organs. Immunol. Rev. 156:25–37. 44. Stolte, M., and S. Eidt. 1989. Lymphoid follicles in antral mucosa: immune response to Campylobacter pylori? J. Clin. Pathol. 42:1269–1271. 45. Tanaka, Y., T. Imai, M. Baba, I. Ishikawa, M. Uehira, H. Nomiyama, and O. Yoshie. 1999. Selective expression of liver and activation-regulated chemokine (LARC) in intestinal epithelium in mice and humans. Eur. J. Immunol. 29:633–642. 46. Uchida, K., K. Okazaki, A. Debrecceni, T. Nishi, H. Iwano, M. Inai, S. Uose, H. Nakase, M. Ohana, C. Oshima, Y. Matsushima, C. Kawanami, H. Hiai, T. Masuda, and T. Chiba. 2001. Analysis of cytokines in the early development of gastric secondary lymphoid follicles in Helicobacter pylori-infected BALB/c mice with neonatal thymectomy. Infect. Immun. 69:6749–6754. 47. Vanbervliet, B., B. Homey, I. Durand, C. Massacrier, S. Ait-Yahia, O. de Bouteiller, A. Vicari, and C. Caux. 2002. Sequential involvement of CCR2 and CCR6 ligands for immature dendritic cell recruitment: possible role at inflamed epithelial surfaces. Eur. J. Immunol. 32:231–242. 48. van Doorn, N. E., F. Namavar, M. Sparrius, J. Stoof, E. P. van Rees, L. J. van Doorn, and C. M. Vandenbroucke-Grauls. 1999. Helicobacter pylori-associated gastritis in mice is host and strain specific. Infect. Immun. 67:3040–3046. 49. Watanabe, T., M. Tada, H. Nagai, S. Sasaki, and M. Nakao. 1998. Helicobacter pylori infection induces gastric cancer in Mongolian gerbils. Gastroenterology 115:642–648. 50. Wotherspoon, A. C., C. Ortiz-Hidalgo, M. R. Falzon, and P. G. Isaacson. 1991. Helicobacter pylori-associated gastritis and primary B-cell gastric lymphoma. Lancet 338:1175–1176. 51. Wyatt, J. I., and B. J. Rathbone. 1988. Immune response of the gastric mucosa to Campylobacter pylori. Scand. J. Gastroenterol. Suppl. 142:44–49. 52. Zerbib, F., G. Vialette, R. Cayla, A. Rudelli, P. Sauvet, D. Bechade, P. L. Seurat, and H. Lamouliatte. 1993. Follicular gastritis in adults. Relations with Helicobacter pylori, histological and endoscopic aspects. Gastroenterol. Clin. Biol. 17:529–534.
