Induction of Polyamine Oxidase 1 by Helicobacter pylori Causes Macrophage Apoptosis by

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1 JBC Papers in Press. Published on July 9, 2004 as Manuscript M Induction of Polyamine Oxidase 1 by Helicobacter pylori Causes Macrophage Apoptosis by Hydrogen Peroxide Release and Mitochondrial Membrane Depolarization* Rupesh Chaturvedi, Yulan Cheng, Mohammad Asim, Francoise I. Bussiere, Hangxiu Xu, Alain P. Gobert, Amy Hacker, Robert A. Casero, Jr., and Keith T. Wilson Department of Medicine, Division of Gastroenterology, and Greenebaum Cancer Center, University of Maryland School of Medicine, Baltimore, Maryland, Veterans Affairs Maryland Health Care System, Baltimore, Maryland, 21201, Department of Oncology, Sidney Kimmel Comprehensive Cancer Center, Johns Hopkins University School of Medicine, Baltimore, Maryland To whom correspondence should be addressed: University of Maryland School of Medicine, 22 South Greene St., Room N3W62, Baltimore, MD Phone: ; fax: ; kwilson@umaryland.edu Running Title: PAO1 in H. pylori-induced macrophage apoptosis Copyright 2004 by The American Society for Biochemistry and Molecular Biology, Inc.

2 2 Summary Helicobacter pylori infects the human stomach by escaping the host immune response. One mechanism of bacterial survival and mucosal damage is induction of macrophage apoptosis, which we have reported to be dependent on polyamine synthesis by arginase and ornithine decarboxylase (ODC). During metabolic back-conversion, polyamines are oxidized and release H 2 O 2, which can cause apoptosis by mitochondrial membrane depolarization. We hypothesized that this mechanism is induced by H. pylori in macrophages. Polyamine oxidation can occur by acetylation of spermine or spermidine by spermidine/spermine N 1 -acetyltransferase (SSAT) prior to back-conversion by acetyl polyamine oxidase (APAO), but recently, direct conversion of spermine to spermidine by the human polyamine oxidase h1, also called spermine oxidase, has been demonstrated. H. pylori induced expression and activity of the mouse homologue of this enzyme (PAO1) by 6 h in parallel with ODC, consistent with the onset of apoptosis, while SSAT activity was delayed until 18 h when late stage apoptosis had already peaked. Inhibition of PAO1 by MDL or by PAO1 sirna significantly attenuated H. pylori-induced apoptosis. Inhibition of PAO1 also significantly reduced H 2 O 2 generation, mitochondrial membrane depolarization, cytochrome c release, and caspase-3 activation. Overexpression of PAO1 by transient transfection induced macrophage apoptosis. The importance of H 2 O 2 was confirmed by inhibition of apoptosis with catalase. These studies demonstrate a new mechanism for pathogeninduced oxidative stress in macrophages, in which activation of PAO1 leads to H 2 O 2 release and apoptosis by a mitochondrial-dependent cell death pathway, contributing to deficiencies in host defense in diseases such as H. pylori infection.

3 3 Introduction Helicobacter pylori is a Gram-negative, microaerophilic bacterium, which selectively colonizes the mammalian stomach, and causes gastritis, peptic ulcers, and gastric cancer. Intriguingly, the human host mounts a vigorous innate and adaptive immune response, yet this results only in lifelong gastritis without eradication of the organism. H. pylori has evolved several strategies to enhance its own survival in the face of this immune response. For example, we have shown that while the host produces nitric oxide (NO) 1 derived from inducible nitric oxide synthase in response to soluble products of H. pylori (1, 2), an arginase enzyme expressed by the bacterium can competitively inhibit host NO production and prevent NO-mediated killing (3). Additionally, H. pylori has been shown to induce apoptosis of T cells (4) and macrophages (5), which is likely to contribute significantly to the persistence of the infection by resulting in immune evasion. In the macrophage studies (5) we demonstrated that H. pylori increased expression and activity of the enzyme arginase II, which produces L-ornithine from L-arginine, and increased activity of ornithine decarboxylase (ODC), which produces putrescine from L- ornithine, that is then converted to the polyamines spermidine and spermine. H. pylori-induced macrophage apoptosis was NO-independent, but was abrogated by inhibition of either arginase or ODC, and restored by addition of spermidine or spermine (5). These studies raised the question as to the mechanism of polyamine-driven macrophage apoptosis. While ODC is the rate-limiting enzyme for polyamine synthesis, there are several pathways of polyamine metabolism that are relevant to their biological function in cells. Spermine and spermidine can be metabolized by an oxidative process that results in the release of H 2 O 2. Specifically, this can occur in two ways. In the originally identified back-conversion pathway, spermine or spermidine are metabolized by the enzyme spermidine/spermine N 1 -

4 4 acetyltransferase (SSAT) to acetylspermine, or acetylspermidine, respectively (6-8), which are then acted upon by N-N peroxisomal polyamine oxidase (PAO) to form spermidine or spermine, respectively (6); this enzyme has recently been cloned (9). Subsequently, this enzyme has been named APAO to denote acetyl PAO (10). More recently, an additional enzyme, PAOh1, was cloned in human cells (11, 12), and the same enzyme was later named spermine oxidase (SMO) (13), since it directly and specifically back-converts spermine to spermidine without the intermediate acetylation step. In the current report, we are studying the mouse homologue, which we will term PAO1. Apoptosis induced by polyamine analogues has been attributed to oxidation of polyamines and generation of H 2 O 2 in cell types that include small cell lung cancer (14), lung adenocarcinoma (15), Chinese Hamster Ovary (16), and breast cancer (17). Release of H 2 O 2 has been linked to depolarization of mitochondrial membrane potential (Δψ m ; (18-20), which is strongly associated with the development of apoptosis by release of cytochrome c from mitochondria to the cytosol (19, 21, 22) and activation of caspases (20, 23). We hypothesized that oxidation of polyamines generated by H. pylori was the key step in causing macrophage apoptosis. We therefore sought to determine the respective roles of the SSAT-APAO versus the PAO1 pathway, the ability of H. pylori to induce these enzymes, and the relationship to apoptosis. This is the first report of any microbe inducing polyamine oxidation, specifically via induction of PAO1 expression and activity in macrophages. We will show that a cascade of events ensues in which oxidation of spermine results in H 2 O 2 release, mitochondrial membrane depolarization, cytochrome c release, caspase-3 activation, and apoptosis.

5 5 EXPERIMENTAL PROCEDURES Reagents All reagents for cell culture, RNA extraction and RT-PCR were from Life Technologies (Grand Island, NY). MDL 72527, a PAO inhibitor, was a gift from N. Seiler (Strasbourg, France). All other chemicals were purchased from Sigma-Aldrich (St. Louis, MO). Bacteria H. pylori SS1, a mouse-adapted human strain (24) was used. Bacteria were passaged on Brucella agar plates containing 10% sheep blood and were maintained under microaerobic conditions. For experiments, H. pylori were harvested from plates, washed twice, and suspended in PBS. H. pylori lysate (HPL) was prepared with a French press in PBS (1). In initial experiments we determined that HPL induced apoptosis to a similar degree as intact bacteria, so to provide a consistent stimulus as we have in previous studies (1, 25, 26), we used HPL for all of the studies presented herein. Concentrations of bacteria were determined by OD at 600 nm. Cells, and culture conditions The murine macrophage cell line RAW was maintained in DMEM supplemented with 10% FBS, 1% penicillin-streptomycin, 1 mm sodium pyruvate, and 10 mm HEPES at 37 C in a humidified 5% CO 2 atmosphere. For experiments, cells were washed and the same medium without penicillin-streptomycin was added 2 h before stimulation. Multiplicity of infection (MOI) was determined as the ratio of bacteria/eukaryotic cell. In the experiments with HPL, the MOI was based on the number of bacteria prior to lysis (1). Experiments were also conducted with peritoneal macrophagesisolated from wild-type C57BL/6 mice (The Jackson Laboratory, Bar Harbor, ME) as previously described (2, 3, 5). RT-PCR RAW macrophages were seeded at 1 x 10 6 /well in six-well plates. After stimulation, total RNA was isolated using TRIzol reagent. Subsequently, 2 µg RNA from each

6 6 sample was reverse transcribed using 50 U Superscript II reverse transcriptase. PCR was conducted using 2 µl cdna and 1 U Taq DNA polymerase. For ODC, PAO1, APAO, and SSAT 15 pmol each of sense and antisense primers were used with 3 pmol each of β-actin primers in a multiplex reaction (2, 5). One PCR cycle consisted of the following: 94 C for 1 min, 60 C for 1.5 min, and 72 C for 1.8 min. The total cycle numbers were 30 for PAO1 and SSAT, 32 for APAO, and 20 for ODC. A final elongation step of 10 min at 72 C was used for each reaction. Sense and antisense primer sequences and PCR product sizes were as follows: murine PAO1, 5'- CACGTGATTGTGACCGTTTC -3' and 5'-TGGGTAGGTGAGGGTACAGTC-3', 222 bp; murine APAO, 5'-CTTTTCCAGGGGAGACCTTC-3' and 5'-CACACCACCTGGATGAACTG- 3', 250 bp; murine SSAT, 5'-GACCCCTGAAGGACATAGCA-3' and 5'- CCGAAGCACCTCTTCTTTTG-3', 248 bp; murine ODC, 5'-CAGCAGGCTTCTCTTGGAAC- 3' and 5'-CATGCATTTCAGGCAGGTTA-3', 602 bp; and murine/human β-actin, 5'- CCAGAGCAAGAGAGGTATCC-3' and 5'-CTGTGGTGGTGAAGCTGTAG-3', 436 bp. PCR products were run on 2% agarose gels with 0.4 µg/ml ethidium bromide. Stained bands were visualized under UV light and photographed with a digital gel documentation system (EDAS 290 and 1D software; Kodak Digital Science, Rochester, NY). Real-time PCR 2 µg RNA from each sample was reverse transcribed using 50 U Superscript II reverse transcriptase. PCR reactions were performed using an Opticon 2 thermal cycler (MJ Research, Cambridge, MA) and SYBR Green master mix (Molecular Probes, Boston, MA) with 5.4 nm primers for murine PAO1, APAO, SSAT or ODC and β-actin as listed above. Thermal cycling conditions included an initial denaturation step (94 C for 2 min) and 40 cycles (94 C for 30sec; 60 C for 30sec; 72 C for 30 sec). Relative expression was calculated from threshold values (C T ) of target and reference genes.

7 7 Assay for ODC activity ODC activity was determined by a radiometric analysis in which the amount of [ 14 C]O 2 liberated from L-[ 14 C]ornithine was measured as previously described (5). Determination of PAO1 and APAO activity 5 x 10 5 RAW cells in 24-well plates were stimulated with HPL at MOI 100. Enzyme activities were assayed in cell homogenates by a modification of a chemiluminescence assay as described (27). Luminol-dependent chemiluminescence was determined using a Monolight 3010 luminometer with two reagent injectors. Luminol was prepared as a 100 mm stock solution in DMSO and diluted to 100 nm with H 2 O, immediately prior to use. Macrophages were scraped into 500 µl of 80 mm borate buffer, ph 9.0, and homogenized with an Ultra-Turrax (IKA Works, Wilmington, NC). For the assay of PAO1 activity, 100 µl of cell lysate was added to 100 µl of 80 mm borate buffer, ph 9.0, containing 570 mu horseradish peroxidase and 250 µm spermine. All reagents were combined and incubated for 2 min at 37 C, then the tube was transferred to the luminometer. Luminol (5 nmol) was added, and the resulting chemiluminescence was integrated over 20 s. The integral values were calibrated against standards containing known concentrations of H 2 O 2 and the activities were expressed as nmol H 2 O 2 /mg protein/min. The assay was linear in the range of 3-89 nmol H 2 O 2. All of the samples were diluted appropriately to be assayed in the linear range. For APAO activity, cell homogenates were prepared in borate buffer, ph 9.0, exactly as above. 100 µl of the cell lysate were added to 100 µl of 80 mm borate buffer, ph 9.0, containing 570 mu horseradish peroxidase and 250 µm N 1 acetyl spermine (Fluka Chemie, Switzerland) as substrate. This assay mixture was incubated for 2 min at 37 C, and then the tube was transferred to the luminometer. Luminol (5 nmol) was added, and the resulting chemiluminescence was integrated over 20 s. Integral values were calibrated and expressed exactly as for PAO1 above with the same linear range.

8 8 Determination of SSAT activity SSAT activity was determined in RAW cell extracts by an assay that measures the formation of L-[ 14 C]acetylspermidine from L-[ 14 C]acetyl- CoA (NEN, Boston, MA) in 10 min at 30 C as described previously (28). The assay mixture contained 50 mm Tris-HCl (ph 7.8), 3 mm spermidine, and 12.7 µm (specific activity 63 mci/mmol) L-[ 14 C]acetyl-CoA in a total volume of 100 µl. The reaction was stopped by adding 10 µl of 1M NH 2 -OH HCl and boiling for 3 min. The resulting samples were spotted onto P-81 phosphocellulose discs and counted in a liquid scintillation counter. Enzyme activity was expressed as nmol L-[ 14 C]-acetylspermidine formed/min/mg protein. Polyamine measurement Polyamine levels were determined by pre-column dansylation reverse phase high-performance liquid chromatography as previously reported (29). Protein concentrations were determined by the method of Bradford (1). Measurement of apoptosis Apoptosis was measured with three methods. i. Annexin V-FITC staining. RAW cells ( cells/well) were cultured in 24-well plates in the presence of HPL for 6 24 h. In some experiments RAW cells were cultured in the presence or absence of the H 2 O 2 detoxifying enzyme, catalase ( U/ml) or polyethylene glycolated (PEG)-catalase ( U/ml) for 24 h. Apoptosis was assayed using an annexin V-FITC apoptosis detection kit (Oncogene Research Products, San Diego, CA). Cells were washed with PBS and resuspended in binding media and stained with annexin V-FITC. Cells were incubated for 30 min at room temperature in the dark and counterstained with 10 µl of propidium iodide (PI; 30 µg/ml). 1 x 10 4 cells were analyzed with a flow cytometer (FACScalibur, Becton Dickinson, San Jose, CA). Spectral overlap was electronically compensated using single color control cells stained with PI or FITC. Analysis of the multivariate data was performed with CELLQuest TM software (Becton Dickinson). The upper

9 9 right (Annexin V + /PI + ) quadrant represents late apoptotic cells, and the lower right (Annexin V + /PI ) represents early apoptotic cells, while the upper left (Annexin V /PI + ) and lower left (Annexin V /PI ) quadrants represent necrotic and viable cell populations, respectively. ii. Apoptosis by DNA histogram analysis. RAW cells or peritoneal macrophages ( cells/well in 24-well plates) were stimulated with HPL for 24 h in the presence and absence of MDL 72527, transfected PAO1 sirna or scrambled sirna, or transfected PAO1 cdna. Both adherent and floating cells were collected and fixed in 70% ethanol. Low molecular weight DNA was extracted with 0.2 M phosphate citrate buffer (ph 7.8) and the remaining cells were stained with PI (30), using a cell cycle analysis kit (Roche Molecular Biochemicals, Indianapolis, IN). Cells were analyzed by flow cytometry. 1.5 x 10 5 cells were acquired and analyzed with ModFit LT TM software (Becton Dickinson) using the sub-g 0 /G 1 peak as the apoptotic population (31). iii. DNA nick end labeling of cells. Macrophages (5 x 10 4 /well) were cultured in 4-well plastic chambered slides (Nunc, Naperville, IL), with or without HPL (MOI of 30) and inhibitors, for 24 h. Cells were then washed with PBS, and fixed with 4% formaldehyde. DNA fragmentation was analyzed by TUNEL assay as described (5) using an In Situ Apoptosis Detection Kit (Trevigen, Gaithersburg, MD). The percentage of apoptotic cells was determined after counting of 10 microscope fields at a magnification of 400X. Determination of cell viability Viability of RAW cells was estimated by a colorimetric assay with the Cell Proliferation Kit II (Roche). The tetrazolium salt 2,3-bis[2- methoxy-4-nitro-5-sulphophenyl]-2h-tetrazolium-5-carboxyaniline (XTT), which is metabolized to formazan by intact mitochondrial dehydrogenases, and electron coupling reagent were added after 20 h and left in the medium for the remaining 4 h of the stimulation period. The viability of cells was estimated on the basis of formazan formed, which was detected spectrophotometrically.

10 10 Transient Transfection of PAO1 sirna in macrophages sirna duplexes were utilized that targeted mouse PAO1 nucleotides 467 to 487, numbered from the start codon (sense, 5'- GGACGUGGUUGAGGAAUUC-3'; antisense, 5'-CCUGCACCAACUCCUUAAG-3'). Scrambled control sirna that has no sequence homology to any known genes was used as the control. 20 µl of the 20 µm stock duplex sipao1 or control scrambled sirna were mixed with 100 µl of optimem medium (Invitrogen, Carlsbad, CA). This mixture was gently added to a solution containing 5 µl of Lipofectamine 2000 (Invitrogen) in 100 µl of optimem. The solution was incubated for 30 min at room temperature and gently overlaid onto 90% confluent RAW cells in 1 ml of optimem for 18 h. Media was changed and cells were incubated for 6 h in DMEM media. Cells were treated with HPL for 6 24 h. After 6 h RNA was isolated and PAO1 mrna expression assessed by RT-PCR using primers that flank the target sequence as follows: sense, 5'-CAATGGCCTTTTGGAAGAGA-3' and antisense, 5'- TTACCATGCCGGAAGAACTC-3'. Apoptosis and PAO1 enzyme activity was performed after 24 h. Transient Transfection of PAO1 in macrophages RAW cells were cotransfected with 400 ng psv-β-galactosidase and 200 ng of pcdna3.1-pao1 using Lipofectamine PLUS (Life Technologies) and optimem media. Cell culture medium was changed 6 h after transfection to complete DMEM medium and cells were stimulated with HPL for 24 h. Transfected cells were stained with PI and analyzed for apoptosis and PAO1 activity as described above. Transfection efficiency was calculated by measuring the β-galactosidase activity in transfected cells. Flow Cytometric Detection of H 2 O 2 by CM-H 2 DCFDA To measure intracellular H 2 O 2 we used the cell permeable redox-sensitive dye CM-H 2 DCFDA (Molecular Probes, Eugene, OR); the nonfluorescent reduced form is converted to the fluorescent form when oxidized, allowing

11 11 detection by flow cytometry (14, 32, 33). CM-H 2 DCFDA is oxidized by cellular H 2 O 2, hydroxyl radicals, and other free radical products of H 2 O 2, while it is relatively insensitive to oxidation by superoxide (32, 33). RAW cells were treated for 1 6 h with HPL, with or without MDL 72527, catalase, or PEG-catalase, washed with PBS, and treated with 10 µm CM-H 2 DCFDA for 20 min at 37 C. 1 x 10 5 cells were analyzed on a Becton Dickinson FACScan for changes in fluorescence (14). H 2 O 2 measurement with Amplex Red 5 x 10 5 cells were plated in 24 well plates. Cells were stimulated with HPL for 6 h, washed with cold PBS, and incubated with a reaction mixture containing 50 µm Amplex Red reagent (Molecular Probes) and 0.1 U/ml horseradish peroxidase in Krebs-Ringer phosphate buffer for 10 min. Plates were read with a fluorescent microplate reader with excitation of 530 nm and emission detection at 590 nm, and H 2 O 2 level determined by using a standard curve with varying dilutions of H 2 O 2 (34). Mitochondrial membrane potential The electron gradient across the mitochondrial membrane space during normal respiration is the mitochondrial transmembrane potential (Δψ m ). Loss of Δψ m (depolarization) was measured by flow cytometry after staining with a MitoCapture TM kit (Calbiochem, San Diego, CA) according to the manufacturer s protocol. The cationic dye fluoresces red as it aggregates inside healthy mitochondria. In apoptotic cells, if the Δψ m collapses, the dye stays as a monomer in the cytoplasm and emits green light. Immunocytochemistry for cytochrome c RAW cells were plated on slide chambers (1 x 10 4 cells/well) and stimulated with HPL, with or without MDL for 18 h. Cells were fixed with 4% paraformaldehyde for 15 min and permeabilized with 100% chilled methanol for 4 min on ice. Slides were blocked with 5% goat serum in PBS for 60 min at room temperature, and then incubated with a monoclonal anti-mouse cytochrome c antibody (Oncogene) overnight at

12 12 4 C. A rabbit anti-mouse secondary antibody conjugated to FITC was used, and staining was visualized with a Nikon ECLIPSE E 800 microscope. Immunoblotting for cytochrome c RAW cells were treated with HPL with or without MDL and incubated for 18 h. Mitochondrial and cytoplasmic fractions were prepared with a cytosolic/mitrochondrial fractionation kit (Oncogene). Protein concentration was determined by the method of Bradford (1) and cytosolic and mitochondrial fraction proteins (40 µg/well) were separated on 16% SDS tris-hcl gels and transferred to PVDF membranes (Bio- Rad, Hercules, CA) by semidry electrotransfer. Membranes were blocked for 2 h at room temperature with PBS containing 0.1% Tween and 5% nonfat dry milk and incubated overnight with anti-cytochrome c antibody at 1:200 dilution. This was followed by a rabbit anti-mouse polyclonal antibody conjugated to horseradish peroxidase (1:5000 dilution). Chemiluminescent detection was performed using the SuperSignal West Pico Chemiluminescent Substrate (Pierce, Rockford, IL) and exposure to Hyperfilm ECL (Amersham, Little Chalfont, U.K.). Measurement of caspase-3 activity Caspase-3 activity was measured by the cleavage of the chromogenic tetrapeptide (AcDEVD-pNA) using a kit from Calbiochem. In brief, 1 x 10 6 cells were lysed and combined with substrate in the caspase reaction mix and incubated at 37 C in the incubation chamber of a SPECTRAmax PLUS microplate reader (Molecular Devices, Sunnyvale, CA). Absorbance was read at 405 nm for 3 h at intervals of 10 min. The conversion factor for the microplate reader was calculated with 100 µl of 50 µm p-nitroaniline. Caspase-3 activity was expressed as pmol/min/mg protein. Statistical Analysis For comparisons between multiple groups, the Student-Newman- Keuls test was used, and for single comparisons between two groups, Student s t test was used. Statview v (SAS Institute, Cary, NC) for the Macintosh was used.

13 13 RESULTS HPL induces gene expression and enzyme activity of ODC, PAO1, and SSAT, but not APAO in macrophages We used murine RAW macrophages that we have previously used to demonstrate H. pylori-induced apoptosis (5), and gene expression (1-3). Macrophages were stimulated with HPL at MOI of 100 for 6 h and mrna expression assessed by RT-PCR. As shown in Fig. 1A, levels of ODC, PAO1, and SSAT were upregulated, while APAO was not induced. This pattern was confirmed by real-time PCR analysis (Fig. 1B), with fold increases of 11.9 ± 2.8 (p < 0.01), 3.7 ± 0.3 (p < 0.001), and 3.8 ± 0.3 (p < 0.001) for ODC, PAO1, and SSAT, respectively, compared to unstimulated macrophages. In contrast, APAO levels were slightly decreased by 0.8 ± 0.2-fold. Activity of each enzyme was measured 6 24 h after stimulation with HPL. There was a biphasic increase in activity of ODC (Fig. 2A), with a rapid peak at 6 h (5.9 ± 0.3-fold increase) followed by a decline at 12 h, and a second peak at 18 h. PAO1 activity (Fig. 2B) was significantly increased at 6 and 12 h, with 3.9 ± 0.9- and 6.1 ± 0.1-fold increases, respectively, with peak activity at 18 h (11.7 ± 0.2-fold increase). In contrast, SSAT was not increased until 18 h (5.3 ± 1.2-fold increase), and APAO was not induced at any of the time points. To further explore the significance of the H. pylori-induced enzyme activation, we determined the putrescine, spermidine and spermine levels at 6 24 h after HPL activation (Fig. 2C). There was no detectable level of putrescine up to 12 h, but there was a marked increase at 18 h that was then significantly reduced from this level at 24 h. In contrast, there was an increase in spermidine at 6 and 12 h, and an increase in spermine at 12 h, followed by a clear decline in these two polyamines that was inversely proportional to the increase in putrescine from h. These polyamine data are consistent with the back-conversion of spermine to spermidine that is

14 14 mediated by the induction of PAO1 from 6 18 h. The subsequent increase in putrescine is likely due to the increase in SSAT activity and subsequent back-conversion of acetylated polyamines by APAO. Time course for HPL induced macrophage apoptosis correlates with activation of ODC and PAO1 activities Because polyamine synthesis and oxidative catabolism have been implicated in the apoptosis of epithelial cell lines, we compared the time course of ODC and PAO1 activation to that of apoptosis in H. pylori-stimulated macrophages. By using annexin V plus PI labeling of live cells, we measured both early and late apoptosis. Fig. 3A indicates that early apoptosis, representing annexin V + /PI cells (right lower quadrants in density plots of Fig. 3B) was significantly increased by 2.1 ± 0.2-fold at 6 h and by 5.6 ± 0.3-fold at 12 h after HPL stimulation, as compared to control. As the early apoptosis decreased from 12 to 24 h, there was a concomitant increase in late apoptosis that peaked at 18 h, an indication that early apoptosis was truly representative of a progressive apoptotic process. The presence of apoptosis beginning at 6 h is consistent with the initial spike in ODC activity at 6 h and the increase in PAO1 activity at this time point, and argues against an important role of the SSAT-APAO pathway in the apoptosis, since SSAT activity did not increase until 18 h after stimulation (Fig. 2B). Macrophage apoptosis is dependent on polyamine oxidation Because the time course of induction of gene expression and enzyme activity of PAO1 and polyamine back-conversion correlated with apoptosis, we determined the effect of an inhibitor of PAO, MDL (4). To quantify the endpoint of apoptosis we used the sub G 0 /G 1 peak of PI-stained fixed cells analyzed by flow cytometry to measure the apoptosis in these experiments. As shown in Fig. 4A and C, there was a 14.6 ±1.7-fold increase in apoptosis at 24 h after HPL stimulation, and MDL inhibited this apoptosis by 35.5 ± 8.8% at 25 µm and by 83.5 ± 3.7% at 250 µm. To verify that

15 15 the apoptosis analysis correlated with cell death, we measured cell viability (Fig. 4B) and found that HPL reduced macrophage survival by 54.3 ± 1.3%, and MDL restored cell viability in a concentration-dependent manner. Because we had found that the increase in early apoptosis in Fig. 3 paralleled the increase in PAO1 activity in Fig. 2B, we further assessed this correlation by determining the effect of MDL on the early apoptosis between 6 and 24 h after stimulation with HPL. As shown in Fig. 4D, the increase in early apoptosis (annexin V + /Pi cells) at 6 24 h was significantly attenuated by MDL Since SSAT is not induced until 18 h, the inhibition of the apoptosis by PAO inhibition at 6 h and 12 h must be due to inhibition of PAO1. This provides additional evidence that the induction of macrophage apoptosis in response to H. pylori is due to PAO1 rather than SSAT/APAO. We next determined if the H. pylori results in the RAW cell line occurred in nontransformed macrophages. As shown in Fig. 4E, when freshly isolated mouse peritoneal macrophages were stimulated with HPL, there was a 5.7 ± 1.1-fold increase in apoptosis, that was inhibited by 74.5 ± 3.1% with MDL 72527, indicating that the same mechanism of apoptosis is occurring in these cells. Finally, we sought to directly address whether PAO1 is essential to H. pylori-induced macrophage apoptosis. We therefore specifically inhibited PAO1 by transiently transfecting RAW cells with PAO1 sirna and compared results to a scrambled control sirna. As shown in Fig. 5, PAO1 sirna markedly decreased the HPL-stimulated PAO1 mrna expression (Fig. 5A) and completely blocked the induction of PAO1 enzyme activity (Fig. 5B). When apoptosis was assessed (Fig. 5C), the PAO1 sirna completely abolished the HPL-induced apoptosis and restored the cell viability to above unstimulated control levels (Fig. 5D). Taken

16 16 together, these data show that knockdown of stimulated PAO1 expression eliminated HPLinduced macrophage apoptosis. Since the inhibition of apoptosis was complete, we have provided direct evidence that PAO1, specifically, is responsible for the apoptosis. Overexpression of PAO1 induces apoptosis in macrophages To provide further proof of principle concerning the causal role of PAO1 in macrophage apoptosis, we transiently transfected RAW cells with a full-length cdna for PAO1. When compared with cells transfected with vector alone, there was a 7.3 ± 0.9-fold increase in PAO1 activity in transfected cells that was nearly identical to the 7.4 ± 1.3-fold increase with HPL-stimulation of neotransfected cells (Fig. 6A). When apoptosis was assessed by DNA histogram analysis in these cells (Fig. 6B), there was a 7.2 ± 0.5-fold increase in apoptosis with PAO1 transfection that exactly paralleled the increase in enzyme activity, and was comparable to the increase in apoptosis with HPL stimulation of neo-transfected cells. H. pylori-induced PAO1 results in H 2 O 2 generation in macrophages and apoptosis Since we had demonstrated that the time course of activation of PAO1 paralleled that of the onset of apoptosis, inhibition of PAO1 by a pharmacologic inhibitor or by PAO1 sirna markedly attenuated apoptosis, and PAO1 overexpression caused apoptosis, we sought to directly determine if H 2 O 2 released by PAO1 activation was responsible for the apoptosis. First, we measured intracellular H 2 O 2 levels in response to HPL by flow cytometry (Fig. 7) in the presence of MDL or the H 2 O 2 detoxifying enzyme, catalase. We assessed time points from 0 6 h because we had identified that there was significant induction of PAO1 and early apoptosis at 6 h, and later time points may have non-specific release of H 2 O 2 from apoptosis. As shown in Fig. 7A and B, there was significant inhibition of HPL-stimulated H 2 O 2 generation at 4 h and 6 h by MDL Additionally there was significant attenuation of H 2 O 2 generation by catalase at 6 h.

17 17 Because the CM-H 2 DCFDA may also detect other oxyradicals, we performed the Amplex Red assay that specifically measures H 2 O 2 in solution. As shown in Fig. 7C, supernatant levels of H 2 O 2 were increased by 2-fold by HPL stimulation and this increase was inhibited by 96.2 ± 2.4 % by MDL and by 86.9 ± 3.6% by catalase. The increased efficiency of H 2 O 2 reduction by catalase in the extracellular assay compared to the intracellular assay data in Fig. 7A most likely reflects the limited uptake of catalase by cells. To further address this, we have used the cell permeable PEG-catalase and found that there was an 80% inhibition of HPL-stimulated intracellular H 2 O 2 generation in the flow cytometric assay (data not shown). Taken together, these data indicate that polyamine oxidation is a major source of oxidative stress, and H 2 O 2 generation, specifically in H. pylori-stimulated macrophages. To directly implicate the H 2 O 2 generation in the apoptosis, we measured the effect of catalase on HPL-stimulated apoptosis. To avoid the confounding effects of H 2 O 2 released nonspecifically during the late phase of apoptosis, we assayed apoptosis with annexin V and PI in live cells, as in Fig. 3, and used MOI of 30. As shown in Fig. 8A and B, catalase inhibited apoptosis in a concentration- dependent manner with 24.9 ± 7.0%, 52.6 ± 4.0%, and 83.5 ± 3.1% inhibition for 250, 500, and 1000 U/ml, respectively. We also assessed apoptosis in the presence of the cell permeable PEG-catalase, and found that there was again a concentration-dependent inhibition of apoptosis, with 100% inhibition at 250 U/ml. These studies indicate that H 2 O 2 generation is a major cause of HPL-induced apoptosis. HPL-induced PAO1 activity results in mitochondrial membrane depolarization, cytochrome c release from mitochondria to cytosol, and caspase-3 activation Because depolarization of Δψ m has been implicated in apoptosis, we determined the ability of H. pylori to cause this event. As shown in Fig. 9A, there was a significant decrease in Δψ m at 12 h, that

18 18 peaked at 18 h. Depolarization of Δψ m was significantly inhibited by MDL (by %) at the time points from h (Fig. 9A). To confirm activation of the mitochondrial apoptosis pathway, release of cytochrome c from mitochondria to cytosol was assessed by immunoblotting. As shown in Fig. 9B, with HPL stimulation, there was a significant decrease in mitochondrial cytochrome c, and a concomitant increase in cytoplasmic cytochrome c, indicating translocation of cytochrome c from mitochondria to cytosol. In cells treated with MDL 72527, there was inhibition of both the decrease in mitochondrial levels and the increase in cytosolic levels of cytochrome c, indicating the prevention of the translocation of cytochrome c. These findings were confirmed by immunohistochemistry. As shown in Fig. 9C, unstimulated cells exhibited a pattern of punctate staining of perinuclear organelles (mitochondria), while HPL activation resulted in intense and diffuse cytosolic staining for cytochrome c (Fig. 9D). Activation in the presence of MDL (Fig. 9E) resulted in an appearance similar to the control cells with a restoration of punctate staining of perinuclear organelles indicating inhibition of cytochrome c release from mitochondria to cytosol. Cytochrome c released from mitochondria activates caspase-9, which ultimately activates caspase-3, a final step in activation of exonucleases and apoptosis (35). Upon activation with HPL, macrophage caspase-3 activity increased significantly by 8.3 ± 2.3-fold and 13.5 ± 1.5-fold at 18 and 24 h, respectively (Fig 9F). As shown in Fig. 9G, MDL significantly reduced caspase-3 activity by 86.0 ± 5.6%. Confirmation by TUNEL assay that polyamine oxidation regulates H. pylori-stimulated apoptosis To directly visualize apoptosis, we performed TUNEL staining (Fig. 10). There was a significant increase in cellular changes consistent with apoptosis with HPL stimulation (Fig. 10B) when compared with the rare apoptotic cells in the unstimulated macrophages (Fig. 10A). In the presence of MDL (Fig. 10C) or catalase (Fig. 10D), there was a marked reduction

19 19 in apoptosis of 68.8 ± 4.0% and 74.1 ± 5.7%, respectively (Fig. 10E), similar to our previously reported results with DFMO in this assay (5).

20 20 DISCUSSION In our previous work we demonstrated that both arginase and ODC are required for H. pylori-induced macrophage apoptosis (5). Intriguingly, we had identified that addition of spermine or spermidine alone, or to H. pylori-stimulated cells treated with the ODC inhibitor α- difluoromethylornithine (DFMO), caused apoptosis, but addition of putrescine had no such effect. This led us to speculate that a product of spermine or spermidine metabolism was required to cause apoptosis. Although regulation of polyamine catabolism was previously attributed to the inducible enzyme SSAT, the recently cloned, inducible enzyme PAOh1/SMO has now been shown to also be an important regulator of polyamine catabolism (11-13). Our current data directly implicate PAO1 induction by H. pylori and represent the first demonstration of the induction of polyamine oxidation as an important component of microbial pathogenesis. Further, we now demonstrate that induction of ODC alone is not sufficient to induce apoptosis in H. pylori-stimulated macrophages and that PAO1 activation is required. H. pylori is considered a non-invasive organism because it lives in the mucus layer of the stomach, where it can adhere to gastric epithelial cells and induce cytoskeletal rearrangements and signaling events such as NF-κB activation and pro-inflammatory IL-8 secretion (36). However, H. pylori antigens have been demonstrated in the lamina propria of the sub-epithelial compartment of the gastric mucosa (37). Because we have shown that contact of live bacteria is not required to induce macrophage apoptosis, we utilized H. pylori lysates in the current study to provide a standardized preparation for our studies and to mimic the exposure of mucosal macrophages to H. pylori components rather than intact bacteria. We previously demonstrated induction of ODC activity with H. pylori stimulation at 24 h (5), and we have now conducted time course studies revealing that the activity actually peaks at

21 21 an early time point of 6 h after stimulation. Additionally, by using real-time PCR we demonstrated a 10-fold increase in ODC mrna expression. In preliminary studies using an ODC promoter construct (38) cloned into a luciferase reporter system, we have observed >8-fold increases in ODC promoter activity with H. pylori stimulation at 6 h, and we have also observed that there is a time-dependent, parallel increase in ODC mrna and enzyme activity between 0 and 6 h (data not shown). Thus, there is an early induction of ODC that primes the system for induction of apoptosis by providing substrate for release of H 2 O 2 by the polyamine oxidation that we have described in this report. A similar early induction of ODC has been reported in LPSactivated peritoneal macrophages (39). The role of induction of ODC in apoptosis is controversial, but ODC-mediated apoptosis has been reported in response to factors that include cytotoxic agents, heat shock, and gamma radiation (40, 41). High levels of ODC expression have been shown to induce apoptosis in interleukin-3 (IL-3)-dependent 32D.3 myeloid cells (42) and transient increase in ODC activity in HL-60 cells treated with etoposide has also been linked to apoptosis (43). Persistent high level expression of ODC has been more strongly associated with cellular proliferation, rather than apoptosis in immune cells such as thymoctyes (40) and leukemia cells (44). In our study, ODC activity transiently peaked at 6 h, declined at 12 h and had a second peak at 18 h. It is likely that the biphasic pattern of the ODC activity is related to the variation in the intracellular polyamine levels that we have observed. An important question that arises is why is there an increase in spermine from 6 12 h at the time that PAO1 is increasing. It is important to realize that at 6 h when ODC activity is highest, there is no increase in spermine, but an increase in spermidine, and that the increase in spermine at 6 12 h is small. Both of these findings are indicative of

22 22 back-conversion of spermine to spermidine by PAO1, because PAO1 is significantly increased at both 6 and 12 h above control levels, while SSAT is not increased until 18 h. An additional issue that we addressed was the relative importance of APAO versus PAO1 in H. pylori-induced polyamine-mediated apoptosis. Our data clearly show that PAO1 mrna levels were increased by 6 h and there is a corresponding increase in enzyme activity at 6 and 12 h. Recently cloned splice variants of the human PAOh1 gene, now recognized as PAO1, have been found to be inducible by specific polyamine analogues in lung cancer cell lines (11, 15). Similarly, increased PAO1 mrna expression has been reported in kidney ischemia-reperfusion injury (45). In contrast, neither APAO mrna levels nor activity were induced by H. pylori, and while SSAT mrna was upregulated at 6 h, the enzyme activity was not increased until 18 h, suggesting a level of posttranscriptional inhibition. Super-induction of SSAT (>1000-fold increases in activity) by anti-tumor polyamine analogues has been shown to be cytotoxic in epithelial cell lines, and inhibition of SSAT with sirna to 100-fold increases was recently shown to be sufficient to prevent apoptosis (46). Therefore, the 5-fold increase in SSAT we observed with H. pylori-stimulation in the absence of APAO induction is less likely to have a causative role in the apoptosis than the induction of PAO1 activity. When enzymatic activities were compared in macrophage lysates, PAO1 produced > 50-fold more H 2 O 2 than SSAT-APAO. Furthermore, SSAT activation in the range we observed is more likely to be a protective mechanism in cells producing acetyl derivatives for export or for recycling when polyamines are present in excess (47). Although APAO is generally considered a constitutively expressed enzyme whose activity is rate-limited by the availability of its acetylated substrate, it can be induced by specific polyamine analogues (9). However, in the current studies, APAO was not upregulated by H. pylori. Our data showing that MDL significantly inhibited early

23 23 apoptosis at 6 and 12 h after stimulation, when PAO1 activity is increased but SSAT-APAO is not, provides additional evidence that PAO1 mediates the induction of macrophage apoptosis by H. pylori. An issue in our studies is that MDL is a specific inhibitor of polyamine oxidases, but not selective for PAO1. It was originally utilized as an inhibitor of the form of PAO now termed APAO for it is preference for acetylated polyamines as substrate (48), but it has also been shown to inhibit PAO1 (27). In HEK293 cells transfected with PAO1, MDL at 200 µm has been reported to inhibit oxidation of spermine by 44-61% (9, 13) and acetyl-spermine by 84% (9). We interpret our findings of 35% inhibition of apoptosis at 25 µm and 75-83% inhibition at 250 µm to be consistent with the incomplete inhibition of PAO1 that has been reported with MDL in cell lines. We addressed the lack of specificity of MDL by conducting studies with an sirna that effectively knocked down PAO1 mrna expression and enzyme activity, and found that the sirna completely prevented H. pylori-stimulated apoptosis. Therefore, we conclude that PAO1, rather than SSAT-APAO, plays the major role in H. pyloriinduced apoptosis. Further support for the role of PAO1 in apoptosis comes from our finding of induction of apoptosis with transfection of PAO1, indicating that PAO1 alone is sufficient to induce apoptosis in macrophages in the presence of sufficient spermine substrate. Intriguingly, it has been reported that transfection of PAO1 increases ODC activity, but transfection of APAO does not have this effect (9). It is possible that induction of PAO1 by H. pylori contributes to the activation of ODC, potentially by inhibition of negative feedback of spermine on ODC. Ultimately, this interaction contributes to the H 2 O 2 generation and apoptosis. We used a flow cytometric assay using CM-H2DCFDA dye to measure intracellular H 2 O 2. Because H 2 O 2 is also produced by dismutation of superoxide (32) it is possible that we

24 24 could have measured other ROS by this technique. Therefore, we also used the Amplex Red assay, which is highly specific for the detection of H 2 O 2 in the supernatant, to confirm our observations with the CM-H 2 DCFDA. The fact that MDL blocked the H 2 O 2 production in both assays strongly suggests that the major source of H 2 O 2 is from polyamine oxidation, and not from other sources. Similar to these findings, MDL has been shown to inhibit H 2 O 2 generation in NCI H157 cells treated with polyamine analogues (14). Our results demonstrate that H. pylori-induced apoptosis was attenuated with catalase, and abolished with the cell permeable PEG-catalase. Exogenous catalase has been shown to inhibit ceramide-induced ROS generation and apoptosis in RAW cells (49). Consistent with findings that H 2 O 2 generated by polyamine oxidation induced by polyamine analogues can depolarize mitochondria (18), H. pylori induced a significant depolarization of macrophage mitochondria. Translocation of cytochrome c into the cytosol from the mitochondria has been implicated in apoptosis because of its ability to activate the caspase cascade by binding to Apaf-1 (50). H. pylori induced translocation of cytochrome c from mitochondria to cytosol that was blocked by MDL72527, implicating this event in polyamine oxidation-induced apoptosis. Since bacteria are known to produce polyamines and serum contains amine oxidases that could be capable of oxidizing polyamines, it is conceivable that H. pylori-derived polyamines could themselves contribute to the apoptosis we have observed. However, we have measured putrescine, spermidine, and spermine levels from equivalent amounts of H. pylori as used in the present studies for stimulation of macrophages, and found no detectable polyamine levels. Additionally, we have reported that H. pylori readily induces apoptosis in macrophages in the absence of serum (5). Together, these points indicate that macrophage apoptosis is due to oxidation of polyamines generated by the host cells and not the stimulating bacteria. Our

25 25 findings are not limited to the murine system, since we have observed that H. pylori induces loss of cell viability in human U937 monocytes that is blocked by MDL (data not shown). Additionally, our findings may be specific to H. pylori, since we have found that the Gramnegative enteric pathogen Citrobacter rodentium, which causes colitis in mice (51), stimulates apoptosis in intestinal epithelial cells and macrophages in vitro that is not attenuated by MDL (data not shown). The persistence of H. pylori in the human stomach for the life of the host, and the chronic gastritis and risk for gastric cancer that ensues all stems from an ineffective immune and inflammatory response in the mucosa. Our current studies provide new insight into the apoptosis of macrophages that contributes to the dysregulated immune response. Specifically, we have shown that polyamine oxidation by the induction of PAO1 mediates the macrophage apoptosis. We have also observed that this same mechanism of apoptosis occurs in gastric epithelial cells (H. Xu, et al. manuscript in preparation) but not in splenocytes or Jurkat T cells (unpublished observations). We suggest that the activation of ODC and PAO1 is part of the innate immune response to H. pylori. Generation of polyamines can have important effects on the regulation of immune response (52) and in gastrointestinal epithelial cells they have beneficial effects in wound repair (53), but our present report provides new evidence that upregulation of polyamine oxidation is likely to have deleterious effects in mucosal host defense in the setting of exposure to selected pathogens, and perhaps, other triggers of PAO1 activation. It will be important to determine the role of ODC and PAO1 in vivo, and to that end we have preliminary evidence that the expression of both enzymes is upregulated in H. pylori gastritis tissues. Interference with polyamine oxidation may ultimately prove to be a useful strategy in augmenting host immune responses.

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30 Higgins, L. M., Frankel, G., Douce, G., Dougan, G., and MacDonald, T. T. (1999) Infect. Immun. 67, Zhang, M., Caragine, T., Wang, H., Cohen, P. S., Botchkina, G., Soda, K., Bianchi, M., Ulrich, P., Cerami, A., Sherry, B., and Tracey, K. J. (1997) J. Exp. Med. 185, Rao, J. N., Li, J., Li, L., Bass, B. L., and Wang, J. Y. (1999) Am. J. Physiol. 277, G FOOTNOTES *This work was supported by grants from the NIH (DK53620 and DK63626 to KTW and CA51085 and CA98454 to RAC), the Office of Medical Research, Department of Veterans of Affairs (to KTW), and the Crohn s & Colitis Foundation of America (to KTW). Current address: Unité de Microbiologie, INRA de Clermont-Ferrand-Theix, Saint- Genès-Champanelle, France. 1 Abbreviations used in this paper: NO, nitric oxide; ODC, ornithine decarboxylase; SSAT, spermidine/spermine N 1 -acetyltransferase; PAO, polyamine oxidase; APAO, acetyl PAO, SMO, spermine oxidase; DFMO, α-difluoromethylornithine; HPL, H. pylori lysate; MOI, multiplicity of infection; PI, propidium iodide; ROS, reactive oxygen species; Δψ m, mitochondrial membrane potential; PEG, polyethylene glycol.

31 31 Figure Legends FIG 1. H. pylori induces mrna expression of ODC, PAO1 and SSAT in RAW macrophages. Cells were stimulated with French press lysate of H. pylori (HPL) at MOI 100 for 6 h. A, RT-PCR for ODC (602 bp product), PAO1 (222 bp product), APAO (250 bp product), SSAT (248 bp product), and β-actin (436 bp product) is shown. Data are representative of 3 separate experiments in duplicate. B, Real-time PCR analysis, using SYBR Green, data are shown on a logarithmic scale. p < 0.01, *p < versus unstimulated cells. n = 6. FIG. 2. Time course of induction of enzyme activities of ODC, PAO1, SSAT, and associated changes in polyamine levels. RAW macrophages were stimulated with HPL at MOI of 100. In A, ODC activity was measured by conversion of [ 14 C]L-ornithine to [ 14 C]O 2. In B, PAO1 and APAO activities were measured by the luminol method, assessing liberation of H 2 O 2 from either spermine in the case of PAO1, or N 1 -acetylspermine, for APAO. SSAT activity was determined by incorporation of L-[ 14 C]acetylCoA into spermidine and formation of L- [ 14 C]acetylspermidine. In C, polyamine levels were determined by HPLC from macrophage lysates, n = 3 experiments in duplicate for A-C. *p < 0.05, p < 0.01 versus time 0, p < 0.05, p < 0.01 versus peak level. FIG. 3. Time course of early and late apoptosis. RAW cells were stimulated with HPL at MOI of 100. Live cells were stained with annexin V (FITC-labeled) and propidium iodide (PI) and flow cytometry performed. A, summary data demonstrating peak early apoptosis (annexin V positive only) at 12 h, and peak late apoptosis (both annexin V and PI positive) at 24 h. p < 0.01 versus time 0; p < 0.01 versus peak level of early apoptosis. n = 4 experiments

32 32 in duplicate. B, representative density plots of annexin V-FITC versus propidium iodide. The values for percent positive cells in the upper right and lower right quadrants are shown. FIG. 4. Inhibition of H. pylori-induced apoptosis in macrophages by the PAO inhibitor MDL A, summary data of apoptosis in RAW cells stimulated with HPL, determined by PI staining of fixed cells followed by flow cytometry and quantification of apoptosis by ModFit-LT software. B, demonstration that HPL reduces cell viability, determined by XTT assay, with HPL, and inhibition of this effect with MDL in concordance with data in A. C, representative histogram plots of PI stained cells (data summarized in A) showing the increase in cells in the sub-g 0 /G 1 fraction, indicative of apoptosis, with HPL stimulation, and inhibition with MDL D, MDL inhibits HPL-stimulated early apoptosis, as determined by Annexin V + /PI staining by flow cytometry of live cells, as in Fig. 3. E, HPL induces apoptosis, determined by PI staining of fixed cells and flow cytometry (as in A), in mouse peritoneal macrophages, which is inhibited with MDL MOI was 30 in A-C, and 100 in D and E. Concentration of MDL in C-E was 250 µm. For A, B, D, and E, p < 0.01 versus unstimulated cells, p < 0.01 versus HPL only. n = 4 experiments in duplicate for A and E, and 3 in duplicate for B and D. FIG. 5. Transfection with PAO1 sirna inhibits H. pylori-stimulated mrna expression, enzyme activity, and apoptosis in macrophages. RAW cells were transiently transfected with duplex sirna targeted against nt in the coding sequence for murine PAO1 or with scrambled sirna control. A, RT-PCR analysis of cells transfected as indicated, in the absence and presence of HPL for 6 h; PAO1 product size was 189 bp. B, PAO1

33 33 enzyme activity measured as in Fig. 2B, after 24 h; C, apoptosis, measured by PI staining and flow cytometry of fixed cells; D, cell viability, measured by XTT assay. MOI of 100 was used in these studies. p < 0.01 versus scrambled sirna alone without HPL; p < 0.01 for PAO1 sirna versus scrambled sirna. n = 4. FIG. 6. Transfection of PAO1 induces apoptosis in macrophages. RAW cells were transfected with mouse PAO1 in the pcdna3.1 vector. MOI of 100 was used. A, PAO1 enzyme activity, determined by the luminol assay. B, apoptosis, determined by PI staining and flow cytometry. In A and B, transfection is cells transfected with empty vector; p < 0.01 versus transfection control without HPL. n = 4 for A, and n = 6 for B. FIG. 7. Effect of H. pylori on hydrogen peroxide production in RAW macrophages, and inhibitory action of MDL 72527, and catalase. Cells were stimulated with HPL at MOI 30 and live macrophages were treated with CM-H 2 DCFDA, and fluorescence measured by flow cytometry at the time points indicated after stimulation. A, summary of data, p < 0.01 versus time 0; p < 0.01 vs. HPL alone, n = 5 experiments. B, representative fluorescence data. The thick line represents the fluorescence tracing from unstimulated cells. The thin line represents the fluorescence tracing from the cells stimulated with HPL alone or with HPL plus MDL or catalase. Note the shift of the curve to the right with HPL in B, indicating more fluorescence intensity, which is prevented by MDL or catalase. C, Amplex Red assay for H 2 O 2 ; cells were stimulated with HPL at MOI 100 for 6 h, and H 2 O 2 released into supernatants in 10 min was measured. p < 0.01 versus unstimulated control; p

34 34 < 0.01 vs. HPL alone, n = 4 experiments. Concentrations were: MDL (250 µm) and catalase (Cat; 1000 U/ml). FIG. 8. Catalase inhibits H. pylori-stimulated macrophage apoptosis. RAW macrophages were stimulated with HPL at MOI of 30, and apoptosis assessed at 24 h by flow cytometry. Live cells were stained with propidium iodide (PI) and annexin V, and apoptotic cells counted as those positive for both. A, summary data, n = 4 experiments in duplicate. Catalase was added at the time of stimulation with HPL at the concentrations indicated. B, representative data, fluorescence from annexin V staining (FITC-labeled) plotted versus PI fluorescence. Conditions are as marked above the density plots. The percent positive in the lower right and upper right quadrants are shown as in Fig. 3. C, PEG-catalase was added at the concentrations shown to RAW cells stimulated with HPL at MOI 100, n = 4. In A and C, p < 0.01 versus control, p < 0.05, p < 0.01 versus HPL alone. FIG. 9. H. pylori induces mitochondrial membrane depolarization, translocation of cytochrome c from mitochondria to cytosol, and activation of caspase-3 that is inhibited by MDL RAW cells were treated with HPL (MOI 100) in the absence and presence of MDL (250 µm). A, Mitochondrial membrane potential was measured by flow cytometry using MitoCapture TM dye at the times indicated. Summary data of mean relative fluorescence, expressed as negative values to indicate depolarization is shown, n = 3 experiments in duplicate, p < 0.01 versus control, p < 0.01 versus HPL + MDL B, Western blot analysis for cytochrome c (15 kda protein). Upper panel: mitochondrial fraction; middle panel: cytosolic fraction. Equal amounts of protein were loaded per lane (40 µg), and equal loading was verified

35 35 by staining of membranes with Ponceau S and immunoblotting for β-actin. C-E, immunofluorescence photomicrographs (magnification 600X) of macrophages stained with antibody to mouse cytochrome c. C, control; D, HPL (MOI 100); E, HPL + MDL In C, the staining is punctate and focal, consistent with localization to mitochondria. In D, the staining is more intense and diffuse in the cytosol, and in E there is reversal of this pattern toward the appearance in the control. F, cells were treated with HPL (MOI 100) and caspase-3 activity measured by colorimetric assay at the times indicated. G, in cells assessed at 24 h after stimulation, MDL (250 µm) blocked the caspase-3 activation. For F and G, p < 0.01 versus control, p < 0.01 versus HPL, n = 3 experiments in duplicate. FIG. 10. Confirmation of dependence of H. pylori-stimulated apoptosis on polyamine oxidation and H 2 O 2 generation by TUNEL assay. RAW macrophages were treated for 24 h as follows: A, unstimulated control; B, HPL, MOI 30; C, HPL + MDL (250 µm); D, HPL + catalase (1000 U/ml). TUNEL positivity was determined by staining of nuclei with 3,3 - diaminobenzidine (magnification 600X) with methylgreen counterstain. E, summary data, the percentage of apoptotic macrophages represents the mean ± SEM of 10 high power fields (> 500 total cells) in each of two separate experiments. p < 0.01 versus unstimulated cells; p < 0.01 versus HPL-stimulated cells.

36 36 A Ctrl HPL ODC β-actin β-actin PAO1 β-actin APAO B Relative mrna Expression β-actin SSAT Ctrl HPL * * 0.5 PAO1 APAO SSAT ODC Figure 1

37 A ODC Activity (pmol CO 2 /h/mg protein) * B PAO1 and APAO Activities (nmol H 2 O 2 /min/mg protein) C PAO1 APAO SSAT * * SSAT Activity (nmol [ 14 C]acetylspermidine formed /min/mg protein) Put Spd Polyamines (nmol/mg protein) Spm Time (h) Figure 2

38 38 A % Apoptosis B Annexin V Annexin V + PI Time ( h) Control 0 h HPL 6 h HPL 12 h HPL 18 h HPL 24 h Figure 3

39 39 A % Apoptosis (PI Staining) B Cell Viability (% control) HPL MDL (µm) C D % Early Apoptosis (Annexin V + /PI ) HPL MDL (µm) Control HPL HPL + MDL Ctrl * Debris Apoptosis Dip G1 Dip G2 Dip S HPL HPL + MDL Time (h) 50 E % Apoptosis (PI Staining) HPL MDL Figure 4

40 40 A + + HPL Scr PAO1 Scr PAO1 sirna β-actin PAO1 B PAO1 Activity (nmol H 2 O 2 /min/mg protein) C D % Apooptosis (PI Staining) Scrambled sirna Ctrl Ctrl PAO1 sirna HPL HPL Cell Viability (% control) Ctrl HPL Figure 5

41 41 PAO1 Activity (nmol H 2 O 2 /min/mg protein) 1000 A B PAO1 transf HPL % Apoptosis (PI Staining) + PAO1 transf + + HPL Figure 6

42 45 A HPL HPL + MDL Fluorescence Units HPL + Catalase B Time (h) Ctrl HPL Ctrl HPL + MDL Ctrl HPL + Cat C H 2 O 2 (nmol/min/10 6 cells) MDL Cat Figure 7 HPL Inhib

43 A % Apoptosis (Annexin V + /PI ) HPL Cat (U/ml) B Control HPL C HPL + Cat 500 HPL + Cat % Apoptosis (Annexin V + /PI ) HPL + Cat 500 HPL + Cat Figure 7 0 HPL PEG-Cat (U/ml) Figure 8

44 44 A Mitochondrial Membrane Potential (- ψ m ) in Relatvie Flourescence Units Control HPL HPL + MDL Time (h) F 180 G180 Caspase-3 Activity (pmol/min/mg protein) B HPL MDL C D E Mit Cyt c Cytos Cyt c β-actin Time (h) 0 HPL MDL Figure 9

45 45 A B C D E % Apoptosis (TUNEL) HPL Inhibitor MDL Cat Figure 10