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J Bacteriol. Feb 2006; 188(4): 1227–1235.
PMCID: PMC1367249
Oxygen Reactivity of PutA from Helicobacter Species and Proline-Linked Oxidative Stress
Navasona Krishnan and Donald F. Becker*
Department of Biochemistry, Redox Biology Center, University of Nebraska, Lincoln, Nebraska 68588
*Corresponding author. Mailing address: Department of Biochemistry, University of Nebraska, N258 Beadle Center, Lincoln, NE 68588. Phone: (402) 472-9652. Fax: (402) 472-7842. E-mail: dbecker3/at/unl.edu.
Received September 27, 2005; Accepted November 11, 2005.
Proline is converted to glutamate in two successive steps by the proline utilization A (PutA) flavoenzyme in gram-negative bacteria. PutA contains a proline dehydrogenase domain that catalyzes the flavin adenine dinucleotide (FAD)-dependent oxidation of proline to Δ1-pyrroline-5-carboxylate (P5C) and a P5C dehydrogenase domain that catalyzes the NAD+-dependent oxidation of P5C to glutamate. Here, we characterize PutA from Helicobacter hepaticus (PutAHh) and Helicobacter pylori (PutAHp) to provide new insights into proline metabolism in these gastrointestinal pathogens. Both PutAHh and PutAHp lack DNA binding activity, in contrast to PutA from Escherichia coli (PutAEc), which both regulates and catalyzes proline utilization. PutAHh and PutAHp display catalytic activities similar to that of PutAEc but have higher oxygen reactivity. PutAHh and PutAHp exhibit 100-fold-higher turnover numbers (~30 min−1) than PutAEc (<0. 3 min−1) using oxygen as an electron acceptor during catalytic turnover with proline. Consistent with increased oxygen reactivity, PutAHh forms a reversible FAD-sulfite adduct. The significance of increased oxygen reactivity in PutAHh and PutAHp was probed by oxidative stress studies in E. coli. Expression of PutAEc and PutA from Bradyrhizobium japonicum, which exhibit low oxygen reactivity, does not diminish stress survival rates of E. coli cell cultures. In contrast, PutAHp and PutAHh expression dramatically reduces E. coli cell survival and is correlated with relatively lower proline levels and increased hydrogen peroxide formation. The discovery of reduced oxygen species formation by PutA suggests that proline catabolism may influence redox homeostasis in the ecological niches of these Helicobacter species.
All organisms convert proline to glutamate in two successive steps catalyzed by proline dehydrogenase (PRODH) and Δ1-pyrroline-5-carboxylate (P5C) dehydrogenase (P5CDH). In gram-negative bacteria, PRODH and P5CDH activities are combined within the proline utilization A (PutA) flavoenzyme (1, 26, 33, 46, 53, 61). The first step of proline oxidation is performed by the flavin adenine dinucleotide (FAD)-dependent PRODH domain, which couples the 2e oxidation of proline to the reduction of the electron chain transport system of the cytoplasmic membrane. The product of the PRODH reaction, P5C, is subsequently hydrolyzed to γ-glutamic semialdehyde, which is then oxidized to glutamate by the NAD-dependent P5CDH domain of PutA. Glutamate that is formed by the oxidation of proline eventually enters the tricarboxylic acid cycle via α-ketoglutarate. Proline catabolism has been shown to be an important energy pathway in various gram-negative bacteria such as Bradyrhizobium japonicum, Sinorhizobium meliloti, and Helicobacter pylori, with proline serving as a preferred respiratory substrate in various environments (12, 25, 38, 58, 59).
Although the PRODH and P5CDH activities of PutA are conserved in gram-negative bacteria, the regulation of proline utilization is divergent. In certain prokaryotes such as Escherichia coli, Pseudomonas putida, and Salmonella enterica serovar Typhimurium, PutA is an autogenous transcriptional repressor responsible for regulating the expression of the putA and putP (encodes a high-affinity Na+-proline transporter) genes (7, 34, 43, 60). The ability of PutA in these bacteria to switch from a DNA-bound transcriptional repressor to a membrane-bound enzyme is dependent on proline availability and the redox state of the FAD cofactor. In the absence of proline, PutA binds to the intergenic DNA control region of the putA and putP genes to repress transcription (7, 43). Upon proline reduction, PutA translocates from the cytosol to a position on the membrane which results in transcriptional activation of the put genes (37, 42). Thus, put gene expression is regulated by the reduction of the FAD cofactor and subsequent PutA-membrane binding (42, 56, 64, 68). The DNA binding domain of PutA from E. coli has been identified as a ribbon-helix-helix (RHH) motif located at the N terminus (residues 1 to 47) (17). Primary sequence analysis predicts whether PutA proteins from other gram-negative bacteria also contain the RHH domain. Prokaryotes in which PutA lacks transcriptional repressor activity, such as Agrobacterium tumefaciens and Rhodobacter capsulatus, regulate proline utilization via transcriptional factors such as PutR, an Lrp-type activator protein which activates putA gene expression in response to increased proline levels (11, 22, 24).
Besides contributing to cellular energy needs, proline metabolism has been shown to impact intracellular redox homeostasis in a variety of organisms such as fungal pathogens, yeast, and mammalian cells (10, 13, 36). Upregulation of the mitochondrial PRODH enzyme in mammalian cells leads to high proline catabolic flux, increased reactive oxygen species (ROS) production, and eventual cell death via apoptosis (13, 32, 44). Proline also appears to function as an antioxidant, scavenging ROS to minimize intracellular oxidative damage during various abiotic and biotic stresses (10). In yeast, disruption of the PUT1 gene (PRODH) increases oxidative stress tolerance (36). Thus, proline metabolism not only contributes to the energy demands of the cell but also can affect the overall intracellular redox environment (18).
Although the metabolic function of PutA is well known, its role in proline-mediated ROS formation has not been investigated. Here, we explore the impact of proline metabolism in the genus Helicobacter by characterizing PutA enzymes from the murine enterohepatic pathogen Helicobacter hepaticus and the human gastric pathogen H. pylori (52). This work is motivated by two independent studies that previously reported that proline was a preferred energy substrate of H. pylori (38, 58). In fact, 10-fold-higher levels of the amino acid proline were detected in the gastric juice of patients infected with H. pylori relative to noninfected individuals, suggesting that in the gut environment, proline serves as an energy source for opportunistic gastrointestinal pathogens (38). Because of the multifaceted roles of proline, proline not only may serve as an opportunistic source of energy but may also be involved in setting the local redox environment of infection. ROS changes accompany events in gastrointestinal diseases and seem to occur mainly via the host inflammatory response requiring H. pylori to survive an onslaught of ROS during infection (2, 19, 23, 63). However, H. pylori is also known to increase ROS levels in host tissues, leading to epithelial cell damage and increased persistence in the gut (9). H. hepaticus infection also causes oxidative stress in the liver (51). Consequently, proline obtained from host tissue damage may serve as an essential energy source and may support persistent colonization and growth of Helicobacter species (38). In certain environments, PutA could have a critical role in colonization and development of chronic disease that has not been previously recognized.
Primary structure analysis of PutA from H. hepaticus (PutAHh) and PutA from H. pylori (PutAHp) predicts that PutA from these Helicobacter species lacks DNA binding activity and is strictly a bifunctional catabolic enzyme. Unexpectedly, PutAHh and PutAHp exhibit reactivity with molecular oxygen during catalytic turnover with proline that is much higher than that of PutA from E. coli (PutAEc), leading to the formation of reduced oxygen species such as hydrogen peroxide (H2O2). PutAHh and PutAHp appear to have different FAD environments, relative to PutAEc, that facilitate proline oxidase activity. The impact of Helicobacter PutA reactivity with oxygen was assessed by oxidative stress studies in an E. coli model system that demonstrated that the enzyme action of PutAHh and PutAHp is toxic to E. coli.
Chemicals, bacterial strains, and culture conditions.
All chemicals and buffers, unless otherwise noted, were purchased from Fisher Scientific and Sigma-Aldrich, Inc. dl-P5C was synthesized as described previously and quantitated using o-aminobenzaldehyde (o-AB) (35, 54). Restriction endonucleases and T4 DNA ligases were purchased from Fermentas and Promega, respectively. PutAEc and PutA from Bradyrhizobium japonicum (PutABj) were prepared as N-terminal His tag fusion proteins using the vectors PutAEc-pET14b and PutABj-pK8AH as previously described (26, 68). Terrific broth was used for the expression and purification of PutA proteins, while LB and M9 minimal salts media were used for in vivo oxidative stress studies. E. coli strains BL21(DE3)(pLyS) and BL21(DE3) were used for the purification of the recombinant PutA proteins and the oxidative stress studies, respectively. Promoter regions of the putA gene from H. hepaticus and H. pylori were amplified from genomic DNA by PCR using primer sets 5′-GTTTTGATTTTAAGCTAAAATTTGTTTATTTG-3′ and 5′-CACTTGCTCCTGCACTCAGGGCGGAAAC-3′ and 5′-GCAGAGCTTACCTTTTATTTAAGAATTTGGCTT-3′ and 5′-GTGAGTTGTTGCTACAAAATTAAAATTCAAGCG-3′, respectively.
Cloning and purification of PutAHh and PutAHp enzymes.
H. hepaticus strain ATCC 51449 genomic DNA was obtained from Gerald Duhamel, University of Nebraska, Lincoln, Nebr. The putA gene from H. hepaticus was PCR amplified using primers 5′-CAGGGCCATATGATGCAAGAAATTATCCAAGAA-3′ and 5′-GTTAGCAGCCGGATCCTTACTTCAAAACTCTTG-3′, which incorporated NdeI and BamHI sites into the PCR product for subsequent cloning into a pET14b vector. The putA gene from H. pylori was cloned similarly by using H. pylori strain J99 genomic DNA, which was a generous gift from Joseph Barycki, Department of Biochemistry, University of Nebraska, Lincoln, Nebr. The putA gene was PCR amplified using primers 5′-CTTAAATAAAAGGTCATATGATGCAAAAAATCATT-3′ and 5′-CTTATTTAATAGGATCCTTATTTTTCAGCACAGCA-3′, which incorporated restriction NdeI and BamHI sites that were subsequently used to clone the putA gene into pK8AH. The resulting PutAHh-pET14b and PutAHp-pK8AH constructs were confirmed by nucleic acid sequencing of the entire putA genes.
PutAHh and PutAHp were overexpressed as N-terminal His6 and His8 tag fusion proteins, respectively, in E. coli strain BL21(DE3)(pLysS) at 25°C as described previously for PutABj, except that 0.5% Triton X-100 was included during the disruption of the resuspended E. coli cells by sonication at 4°C (26). The PutA proteins were purified using Ni2+-nitrilotriacetic acid and anion-exchange column chromatography as described previously (26, 68). The purity of the protein preparations was >90% as judged by sodium dodecyl sulfate-polyacrylamide gel electrophoresis, and the identity of the PutAHh and PutAHp proteins was confirmed by mass spectral analysis performed in the Metabolomics Core Facility in the Redox Biology Center at the University of Nebraska-Lincoln. The oligomeric properties of PutAHh and PutAHp were characterized by gel filtration chromatography (Superdex-200 column) as described previously (17). PutAHh and PutAHp were stored in 50 mM potassium phosphate (pH 7.5) containing 10% glycerol at −70°C.
Characterization of PutA proteins.
PRODH activity was analyzed using the proline:dichlorophenolindophenol (DCPIP) oxidoreductase assay, and P5CDH activity was determined by the reduction of NAD+ at 340 nm as described previously (6). Proline:DCPIP oxidoreductase assays were performed in 50 mM Tris buffer, pH 7.5, with various amounts of proline (0 to 400 mM). One unit of PRODH activity is the quantity of enzyme that transfers electrons from 1 μmol of proline to DCPIP per min at 25°C. Proline:O2 activity was determined in air-saturated MOPS (morpholinepropanesulfonic acid) buffer (20 mM MOPS, pH 7.5) at 25°C with various amounts of proline (0 to 400 mM). Rates of the proline:O2 activity were determined by following the production of H2O2 or P5C with one unit of activity defined as the amount of PutA that generates 1 μmol of H2O2 or P5C per min at 25°C. H2O2 was quantitated by analyzing aliquots from the oxidase reaction mixture by a catalase colorimetric assay kit (Sigma) using a standard curve of known H2O2 concentrations. The catalase assay was performed as recommended by the manufacturer. Superoxide anion detection during the PutA oxidase reactions was performed as previously described by monitoring cytochrome c reduction at 550 nm in the absence and presence of 30 U of bovine superoxide dismutase (Sigma) (41). For detecting P5C, 4 mM (final concentration) of o-AB was included in the proline:O2 assay mixture, and the formation of the o-AB-P5C yellow complex was monitored at 443 nm (ε = 2,900 M−1 cm−1) (35). All enzyme assays were initiated by adding PutA enzyme and were performed in triplicate. The kinetic parameters Km and kcat were estimated by nonlinear regression analysis of the initial reaction velocity versus proline concentration using the Michaelis-Menten equation. Turnover numbers were calculated using molecular mass values of 135.5 kDa and 137.4 kDa for the monomeric species of PutAHh and PutAHp, respectively. The apparent inhibition constant (Ki) for the inhibition of PutAHh and PutAHp by l-tetrahydro-2-furoic acid (l-THFA) was determined as described previously for PutA from E. coli (69).
The concentrations of PutA proteins were determined using bicinchoninic acid reagents (Pierce) with bovine serum albumin as the standard and spectrophotometrically using molar extinction coefficients at 450 nm of 13,373 ± 281 cm−1 M−1 and 12,944 ± 166 cm−1 M−1 for PutAHh and PutAHp, respectively (6, 62). The molar extinction coefficients for PutAHh and PutAHp at 450 nm were determined as previously described (6). Because it became apparent during this work that purified PutAHh was relatively more stable than PutAHp, further biochemical characterization of PutA from Helicobacter species focused on PutAHh. Proline titrations of PutAHh were performed at 20°C in 50 mM potassium phosphate buffer (pH 7.5) containing 5% glycerol in a glove box (Belle Technology) under a nitrogen atmosphere (26). The protein-proline mixtures were equilibrated 10 min prior to the recording of each spectrum, and the data were analyzed as previously described, assuming the formation of a reduced PutA-P5C equilibrium complex (6). Titrations of PutAHh with sodium sulfite were performed anaerobically under a nitrogen atmosphere in a glove box using sodium sulfite stock concentrations of 2, 20, 200, and 1,000 mM. Spectra were recorded after equilibrating PutAHh with different concentrations of sodium sulfite. The enzyme solution was allowed to equilibrate after each addition of sodium sulfite until no further changes in the absorbance were observed, ~15 to 20 min. To test the reversibility of the FAD-sulfite adduct, l-THFA was added incrementally to the sulfite-PutAHh mixture up to a final concentration of 4 mM l-THFA. After each addition of l-THFA, the PutA solution was equilibrated for 30 min before the spectrum was recorded.
Oxidative stress studies.
Oxidative stress studies were performed in E. coli strain BL21(DE3) in M9 minimal salts medium according to a previously described protocol (16, 67). Cells harboring the putA genes from E. coli, B. japonicum, H. hepaticus, and H. pylori were grown in 20-ml cultures in LB broth at 37°C, and expression of the PutA proteins was induced for 3 h with 0.5 mM isopropyl-β-d-thiogalactopyranoside (IPTG) when the culture reached an optical density of ~0.8. Expression of the PutA proteins was confirmed by PRODH assays of cell extracts from each culture with average values of about 0.33 (PutAHh), 0.43 (PutAHp), 1.2 (PutABj), and 1.2 (PutAEc) U/mg of total protein, indicating an almost fourfold variation in PRODH activity levels. After treatment with IPTG, cell cultures (20 ml) were spun down at 9,000 × g, and the resulting cell pellets were resuspended in 2 ml of M9 minimal salts medium and incubated with various reagents at 37°C for 1 h. H2O2, proline, and the competitive inhibitor l-THFA were added to the 2-ml cultures at a final concentration of 5 mM. After the 1-h incubation, cell cultures were serially diluted and plated onto LB agar plates containing ampicillin (50 μg/ml) and grown overnight at 37°C. The next day (~16 h), colonies were counted and multiplied by the dilution factor to estimate the number of surviving colonies. Results are presented as the percent survival rate, which is calculated as follows: [(number of colonies after addition of reagent)/(number of colonies with no addition of reagent)] × 100. The reported percent survival rates are the average values from at least six independent experiments.
Proline and H2O2 levels were determined in 2-ml cultures supplemented with no proline, 5 mM proline, or a mixture of 5 mM each of proline and l-THFA as described above. For proline measurements, the 2.0-ml cultures in the M9 minimal salt medium were pelleted by centrifugation, resuspended in 0.5 mM of sterile water, and lysed by boiling for 10 min. The lysed samples were then centrifuged for 5 min at 9,000 × g. The resulting supernatant was incubated with 46 mM ninhydrin and 0.2 ml glacial acetic acid for 1 h at 100°C (5). The reactions were stopped by placing the samples on ice. After the mixtures were extracted with toluene (0.4 ml), the absorbance at 520 nm was recorded (5). A proline standard curve ranging from 0 to 3.0 mM proline was used to determine the proline level of each sample. The intracellular proline levels were then estimated by assuming a cytosolic volume of 0.47 μl per ml of culture (optical density, ~1.0) (21). H2O2 in the medium of the cell cultures was monitored by analyzing aliquots at different time points from 0 to 60 min during the incubation as described above. Cells were pelleted by centrifugation, and H2O2 in the supernatant was measured using an Amplex Red hydrogen peroxide/peroxidase assay kit (Molecular Probes). Extracellular H2O2 generated in a control medium of E. coli cells devoid of recombinant PutA was subtracted from the H2O2 measurements of E. coli cells containing different PutA enzymes to correct for nonspecific H2O2 formation. The estimates of proline and H2O2 are average values of at least three independent determinations.
Functional domain analysis and properties of PutAHh and PutAHp.
The PutA enzymes from H. hepaticus (GenBank accession number NC_004917) and H. pylori (GenBank accession number NP_222770) are polypeptides of 1,166 and 1,185 amino acids, respectively, share 63% sequence identity, and have a low sequence identity of 18 to 19% with PutAEc (1,320 amino acids) (3, 55). Domain alignment of Helicobacter PutA enzymes with PutAEc indicates that PutAHh and PutAHp share the PRODH and P5CDH domains but lack the RHH domain of PutAEc. The absence of the RHH domain was confirmed by testing the DNA binding activities of PutAHh and PutAHp by gel mobility shift assays using the corresponding promoter regions of the putA genes from H. hepaticus and H. pylori (6). PutAHh and PutAHp displayed no DNA binding activity even with up to a 100-fold excess of PutA protein compared to DNA (data not shown as negative results). X-ray crystal structures (2.0 to 2.1 Å) of the PutAEc PRODH domain complexed with competitive inhibitors l-lactate and l-THFA have been solved, allowing active-site residues within 5 Å of the noncovalently bound FAD cofactor and the inhibitors to be identified (66). Among the active-site residues in the PRODH domain of PutAEc, 52% are identical in PutAHh and PutAHp, indicating a similar PRODH domain fold. Size exclusion chromatography shows that PutAHh and PutAHp purify as dimer-monomer mixtures with the predominate species being a dimer. PutAEc purifies mainly as dimeric species in which the dimer is stabilized by intermolecular interactions in the RHH domain (17).
The proline:DCPIP oxidoreductase activities of PutAHh and PutAHp are summarized in Table Table1.1. Although slightly higher Km values for proline were observed, similar PRODH turnover numbers relative to PutAEc were determined for PutAHh and PutAHp. The PRODH active sites in PutAHh and PutAHp were explored by evaluating the inhibition of PRODH activity by the competitive inhibitor l-THFA. l-THFA competitively inhibits PutAEc, with a Ki of 0.2 mM (69). PutAHh and PutAHp were also shown to be competitively inhibited by l-THFA, with apparent Ki values of 0.33 and 0.35 mM, respectively, supporting similar substrate binding (data not shown). Specific P5CDH activities in PutAHh and PutAHp were 0.25 and 0.19 U/mg, respectively, which are near the P5CDH specific activity level (0.3 U/mg) for PutAEc (6).
TABLE 1.
TABLE 1.
Kinetic parameters for PutAEc, PutAHh, and PutAHp enzymesa
The UV-visible absorbance spectrum of oxidized PutAHh is shown in Fig. Fig.11 (curve 1), with the main absorbance peaks at 450 and 367. A molar extinction coefficient of 13,373 ± 281 cm−1 M−1 at 450 nm was determined for FAD bound to PutAHh with about 1 mol of FAD bound per polypeptide, similar to PutAEc (6). The same ratio of FAD to polypeptide was found for PutAHp, with a molar extinction coefficient of 12,944 ± 166 cm−1 M−1 at 450 nm. Figure Figure11 also shows a proline titration of PutAHh demonstrating that proline can fully reduce the bound FAD cofactor with an equilibrium constant for the reduction of the enzyme of 14.3 ± 0.43 mM−1 proline. The observation that proline (reduction potential [Em], ~−0.124 V for the proline/P5C couple [pH 7.5]) can fully reduce the FAD cofactor indicates that the reduction potential of the FAD cofactor in PutAHh is more similar to the reduction potential of the FAD cofactor in PutAEc (Em, −0.077 V [pH 7.5]) than to that in PutABj (Em, −0.132 V [pH 7. 5]), which is only partially reduced by the substrate proline (6, 26, 62, 69). Proline-reduced PutAHh is fully reoxidized upon mixing with an equal volume of air-saturated buffer with a half-life of ~12 min. The oxygen reactivity of PutAHh is about 10-fold faster than that of proline-reduced PutAEc, which reacts very slowly with air (half-life, ~120 min) (6). The sluggish oxygen reactivity of proline-reduced PutAEc protects the substrate-derived reducing equivalents until the flavoenzyme reacts with the correct physiological partner, which is presumably ubiquinone, in the electron transport chain.
FIG. 1.
FIG. 1.
Proline reduction of PutAHh. PutAHh was titrated with proline under anaerobic conditions in 50 mM potassium phosphate (pH 7.5) at 25°C. Spectra were recorded 10 min after the addition of proline. Curves 1 to 9, 0, 0.02, 0.04, 0.05, 0.07, 0.37, (more ...)
Proline:O2 activity.
To assess reactivity with molecular oxygen during catalytic turnover with proline, the rates of proline oxidation by PutAHh, PutAHp, and PutAEc were measured by monitoring H2O2 formation in air-saturated buffer. The kinetic parameters are summarized in Table Table11 and show that PutAHh and PutAHp display significant catalytic turnover consistent with the high oxygen reactivity of the proline-reduced enzymes relative to PutAEc. The proline:O2 specific activities for PutAHh and PutAHp correspond to apparent turnover numbers of 29 and 31 min−1, respectively, which are >100-fold higher than the estimated turnover number for PutAEc (<0.3 min−1). The Km values for proline determined for PutAHh and PutAHp are the same in both the proline:DCPIP oxidoreductase and the proline:O2 assays. Evaluation of the kinetic data in Table Table11 also shows that PutAEc has a significant preference for the artificial electron acceptor during catalytic turnover with proline based on the >2,500-fold activity in the proline:DCPIP oxidoreductase assays. In contrast, PutAHh and PutAHp show a more modest preference for the artificial electron acceptor, with about 20-fold-higher activity in the proline:DCPIP oxidoreductase assays than in the proline:O2 assays. The diminished preference for the artificial electron acceptor is due not to lower proline:DCPIP oxidoreductase activity but rather to a noticeable increase in oxygen reactivity. Similar differences between PutAEc and PutAHh in catalytic turnover with proline using oxygen as the electron acceptor were also observed in o-AB chromogenic assays, which monitor the formation of P5C. Turnover numbers of 17 min−1 and <0.3 min−1 were determined for PutAHh and PutAEc, respectively. To test whether the higher proline:O2 activity may be generally associated with bifunctional PutA enzymes, we also evaluated the bifunctional PutABj enzyme, which shares 37% amino acid identity with PutAEc (26). PutABj displayed low proline:O2 activity (1 min−1), similar to PutAEc, indicating that the increased reactivity of PutAHh and PutAHp with molecular oxygen during catalytic turnover with proline is not necessarily a general property of bifunctional PutA enzymes. Formation of a superoxide anion was detected during PutAHh and PutAHp oxidase reactions by cytochrome c reduction, indicating that superoxide is generated during catalytic turnover with proline (data not shown). Superoxide was not detected in assays with PutAEc, consistent with much lower oxygen reactivity.
To further characterize the increased proline:O2 activity of Helicobacter PutA enzymes, we compared the reactivity of the FAD cofactor with sodium sulfite in PutAHh with that in PutAEc. Flavoenzymes that react with molecular oxygen during catalytic turnover typically form reversible sulfite adducts at the N-5 position of the isoalloxazine ring of FAD, which is observed by bleaching of the flavin absorbance at 450 nm (31). Figure Figure22 (top) shows that the addition of sodium sulfite to PutAHh decreases the absorbance of the FAD cofactor at 450 nm. From the change in the absorbance at 450 nm, a dissociation constant of 400 ± 60 μM was estimated for the formation of a FAD-sulfite adduct in PutAHh. In contrast, PutAEc does not react with sulfite even at up to 50 mM concentrations of sodium sulfite (data not shown). The FAD-sulfite adduct was stable in air but was disrupted by the addition of l-THFA to the PutAHh-sulfite mixture. Figure Figure22 (bottom) shows that l-THFA restores the oxidized spectrum of PutAHh, indicating that sulfite is displaced from the PRODH active site. It is evident in Fig. Fig.22 that l-THFA is bound to the PRODH active site, as a prominent shoulder in the FAD absorbance spectrum at 474 nm is observed (69). Thus, PutAHh shows biochemical characteristics that are consistent with higher oxygen reactivity during turnover with substrate.
FIG. 2.
FIG. 2.
Reaction of PutAHh with sodium sulfite. (Top) PutAHh was incubated with different concentrations of sodium sulfite in 50 mM potassium phosphate buffer (pH 7.5) at 25°C for 15 min. Curves 1 to 6, 0, 0.05, 0.1, 0.5, 1, and 4 mM sodium sulfite, respectively. (more ...)
Stress tolerance in Escherichia coli.
To test whether the proline oxidase activities of PutAHh and PutAHp have any impact on redox status in bacteria, we compared the stress tolerance levels of E. coli cultures expressing PutAEc, PutABj, PutAHh, and PutAHp. Figures Figures33 and and44 show the survival of E. coli cells after treatment with 5 mM H2O2. Expression of PutAEc and PutABj, which display minimal oxidase activity, has little effect on the tolerance of E. coli cells to oxidative stress relative to control E. coli cells that contain only a vector (i.e., pET14b). In stark contrast, acute oxidative stress treatment is basically lethal to E. coli cells harboring PutAHh and PutAHp, with survival rates of <0.05%. To explore the toxicity of PutAHh and PutAHp further, E. coli cultures were incubated with 5 mM proline. As described above, PutAEc and PutABj had little effect relative to the control E. coli cells. However, in the presence of proline, expression of PutAHh and PutAHp is toxic to E. coli, with survival rates of around 0.5%. It appears that the enzyme action of PutAHh and PutAHp kills the E. coli cells, perhaps due to the formation of reduced oxygen species. The PRODH activity of PutAHh and PutAHp was then blocked by supplementing E. coli cultures with the competitive inhibitor l-THFA (5 mM) in the presence of proline (5 mM). Figures Figures33 and and44 show that l-THFA rescues E. coli cells expressing PutAHh and PutAHp and increases survival rates to 60 to 70%. Thus, the enzyme action of PutAHh and PutAHp is the causative agent of proline toxicity in E. coli.
FIG. 3.
FIG. 3.
Acute stress treatment of E. coli cell cultures expressing the various PutA enzymes. E. coli cells were incubated with 5 mM H2O2, 5 mM proline, or a mixture of 5 mM each of proline and l-THFA in M9 minimal medium for 1 h at 37°C. After the incubation, (more ...)
FIG. 4.
FIG. 4.
Cell survival rates of E. coli cells expressing different PutA enzymes. E. coli cells were incubated with 5 mM H2O2 (black), 5 mM proline (gray), or a mixture of 5 mM proline and 5 mM l-THFA (diagonal stripes) in M9 minimal medium for 1 h at 37°C. (more ...)
To gain further insights into how the expression of the Helicobacter PutA enzymes affects cell survival, proline and H2O2 levels were measured after E. coli cells were incubated in M9 minimal salts medium for 1 h at 37°C with no supplement, with 5 mM proline, or with a mixture of 5 mM proline and 5 mM l-THFA. Figure Figure55 shows that without proline supplementation, intracellular proline levels are about twofold lower in E. coli cells (2 to 2.3 mM) expressing the PutA enzymes than in control cells (5.2 ± 0.3 mM proline). With proline supplementation in the medium, proline levels were also lower in E. coli cells expressing PutAEc and PutABj (~17 mM) and cells expressing PutAHh and PutAHp (~9 mM) than in control cells (33.2 ± 1 mM). These observations demonstrate that the catabolic activity of recombinant PutA enzymes depletes intracellular proline pools, with larger decreases observed with the Helicobacter PutA enzymes. The estimated intracellular levels of proline are within the range of previous measurements of proline pools in E. coli from studies that evaluated proline uptake and accumulation in response to osmotic stress (4, 39, 65). As expected, the addition of 5 mM l-THFA prevents the breakdown of intracellular proline, as levels in E. coli cells harboring PutA enzymes (27 to 36 mM proline) are in the range of proline content measured for the control cultures (34.5 ± 0.9 mM).
FIG. 5.
FIG. 5.
Proline levels in E. coli cell suspensions expressing different PutA enzymes. E. coli cells were incubated and supplemented with no proline (black), 5 mM proline (gray), or a mixture of 5 mM proline and 5 mM l-THFA (diagonal lines) in M9 minimal medium (more ...)
Next, H2O2 released into the medium from E. coli cells expressing the different recombinant PutA enzymes was measured. Figure Figure6A6A shows that the amount of H2O2 detected in the medium by 60 min was higher in E. coli cells expressing Helicobacter PutA (~1.5 μM) than in cells expressing PutABj (0.32 ± 0.05 μM) and PutAEc (0.28 ± 0.04 μM). Although PutAEc and PutABj decrease the intracellular proline pool, similar to PutAHh and PutAHp (Fig. (Fig.5),5), less H2O2 is generated during proline degradation by these enzymes. Figure Figure6B6B demonstrates that proline supplementation increases H2O2 levels in the external medium of each culture, with estimates at 60 min of 7.3 ± 0.6 μM and 7.2 ± 0.3 μM for PutAHh and PutAHp, respectively. In E. coli cells containing PutABj and PutAEc, H2O2 concentrations of 1.6 ± 0.1 μM and 0.7 ± 0.1 μM were detected, respectively. Thus, the addition of proline to E. coli cells expressing Helicobacter PutA elevates extracellular H2O2 levels nearly 5-fold higher than cells containing PutABj and ~10-fold higher than cells with PutAEc. It is evident from the lower proline levels found in cells harboring PutAHh and PutAHp (Fig. (Fig.5)5) that Helicobacter PutA enzymes create a higher proline catabolic flux than PutAEc and PutABj that is apparently due to increased oxygen reactivity. Figure Figure6C6C shows that the addition of l-THFA restores H2O2 levels, demonstrating that the higher amounts of extracellular H2O2 measured from cells expressing PutAHh and PutAHp are derived from proline oxidation.
FIG. 6.
FIG. 6.
Time course of H2O2 released from E. coli cells harboring different PutA enzymes. Extracellular H2O2 was measured in E. coli cell cultures containing PutAEc ([filled square]), PutABj ([filled lozenge]), PutAHh (•), and PutAHp ([filled triangle]) in M9 minimal medium. (more ...)
PutA enzymes can be classified according to whether they lack (i.e., bifunctional) or are endowed with (i.e., trifunctional) transcriptional regulator activity. Sequence alignments of PutAEc with PutA enzymes from different sources have identified >20 PutA proteins that contain the RHH domain and are thus predicted to function as autogenous transcriptional regulators. For bacteria in which PutA does not function as an autogenous transcriptional regulator, proline utilization requires other regulatory proteins such as PutR and PruR. PutR is an Lrp-type transcriptional repressor that has been identified in R. capsulatus, and PruR is a AraC/XylS family regulator that has been shown to activate expression of the putAP operon in Pseudomonas aeruginosa (24, 40). In the food-borne pathogen Vibrio vulnificus, neither PutR nor PruR appears to be involved; instead, the cyclic AMP receptor protein has been postulated to regulate the putAP genes (28). Genome analysis of H. hepaticus and H. pylori reveals that the putA and putP genes are arranged in an operon similar to those of P. aeruginosa and V. vulnificus. Potential homologues of PutR and PruR in the genomes of H. hepaticus and H. pylori, however, are not apparent, and a cyclic AMP receptor protein binding site (TGTGAN6-8TCACA) is not present in the promoter regions of the putAP operons, suggesting further divergence in mechanisms for the regulation of proline utilization in Helicobacter species.
Surprisingly, PutAHh and PutAHp have significantly higher reactivity with molecular oxygen than PutAEc during catalytic turnover with proline. Accordingly, PutAHh forms a FAD-sulfite adduct, while PutAEc is unreactive, consistent with the notion that flavoproteins that react with sulfite generally have higher oxidase activity. Flavin dehydrogenase enzymes typically react slowly with oxygen during catalytic turnover with substrate to help ensure the efficient transfer of electrons to the correct physiological acceptor for maximum energy utilization of the substrate (29). A notable example includes the mammalian acyl coenzyme A dehydrogenases, which profoundly depress oxygen reactivity in the product-bound form, preventing formation of reactive oxygen species in the mitochondria (14). Similarly, the free-dihydroflavin form of PutAEc reacts rapidly with oxygen but shows significantly lower oxygen reactivity in the substrate/product-bound form (6).
The structural determinants of oxygen reactivity in flavoenzymes are not fully understood, but it seems that the electrostatic environment of the active site is a critical component (30). The first step in the reaction of reduced flavins with molecular oxygen is the formation of a radical pair involving superoxide (O2) and a flavin radical (29). The formation of the superoxide anion is favored by overall positive electrostatic environments, whereas active sites with lower dielectric environments are thought to hinder superoxide formation (14, 15, 30). Structural studies of the conversion of xanthine dehydrogenase to xanthine oxidase indeed show differently charged active sites (15). Xanthine dehydrogenase is a metalloflavoenzyme, which, during catalytic turnover with xanthine, has a preference for NAD+ as an electron acceptor, while the xanthine oxidase form prefers molecular oxygen (20). It is clear from the structural analysis of both forms of the enzyme that the electrostatic environment is more positive in the xanthine oxidase form than in the xanthine dehydrogenase form (15). Kinetic studies of glucose oxidase also suggest that positively charged residues in the active-site environment are important for accelerating oxygen reactivity of FAD cofactors (45). Differences between the electrostatic environments of the PRODH active sites of PutAEc and Helicobacter PutA are not obvious by primary structure analysis alone. PutAHh and PutAHp share 52% of the active-site residues in PutAEc that have been identified by X-ray structure determination of the PutAEc PRODH domain (66). The predicted variations in the active-site residues of PutAHh and PutAHp from PutAEc do not involve changes in charged residues. The only relevant substitution may be that Tyr437 in PutAEc, which is conserved in PutABj as Tyr345, is replaced by Asn292 and Asn291 at the analogous positions in PutAHh and PutAHp, respectively (26, 66). From the structural analysis of the PRODH domain of PutAEc, it appears that Tyr437 separates the active site from bulk solvent (66). Thus, the replacement of Tyr with a smaller side-chain residue such as Asn in Helicobacter PutA may increase solvent accessibility in the active site, which would favor higher oxidase activity. In the mammalian medium-chain acyl coenzyme A dehydrogenase, it has been proposed that product binding desolvates the active site, resulting in lower oxygen reactivity relative to the free enzyme (14). Likewise, proline/P5C binding may more effectively desolvate the active site in PutAEc than in PutAHh or PutAHp. A future goal is to determine the three-dimensional structure of PutA from Helicobacter species to compare the flavin active-site environment and oxygen reactivity with those of PutAEc.
The proline-linked formation of reduced oxygen species by PutAHh and PutAHp was shown to be toxic to E. coli, severely inhibiting cell survival. Blocking PRODH activity rescued cell survival, demonstrating that the enzyme action of Helicobacter PutA is harmful under high-proline conditions. Increased proline catabolism in general, however, was not lethal, as PutAEc and PutABj expression in E. coli had no impact on cell survival despite the detection of increases in H2O2 levels upon the addition of proline to the medium. Therefore, the higher oxidase activity of Helicobacter PutA appears to be the culprit for inducing cell death. Alkyl hydroperoxide reductase, the main scavenger of endogenous H2O2 in E. coli, and catalase are responsible for keeping intracellular H2O2 concentrations below toxic levels (~2 μM) (48-50). Apparently, the H2O2 generated by PutABj and PutAEc does not reach toxic levels, while the higher proline oxidase activity of PutAHh and PutAHp is enough to overwhelm the scavenging system and produce toxic effects under our experimental conditions. Perhaps more importantly, though, is the formation of superoxide during the proline oxidase reaction, which likely has a considerable role in the lethal effect of proline catabolism via Helicobacter PutA and in the acute sensitivity to oxidative stress treatment.
The physiological significance of PutA oxidase activity in Helicobacter species is presently not clear. One simple explanation is that the higher oxidase activity of PutAHh and PutAHp is an outcome of the microaerophilic environment of Helicobacter species, which may not require robust protection from oxygen. On the other hand, PutABj, which is also from a microaerophilic bacterium, resembles PutAEc, with low oxidase activity and no deleterious effect on E. coli cell survival. A potential role of the PutA oxidase activity in Helicobacter species may be that proline metabolism contributes to the redox environment of infection. It has been shown that proline is a preferred respiratory substrate in cultured H. pylori cells and that patients infected with H. pylori have considerably higher levels of proline in the gut (38). Thus, proline not only may be used for energy but may also affect ROS levels which impact colonization and/or the persistence of Helicobacter species in the host. Proline metabolism has been shown to be critical in different ecological niches, with proline having multifaceted roles in protein chaperoning, abiotic stress protection, and energy utilization (8, 10, 18, 27, 36, 38, 47, 52, 57). To address the pathophysiological role of PutA and proline metabolism in Helicobacter species, future work will include the analysis of PutA expression and proline metabolic flux in host infection animal models.
Acknowledgments
This research was supported in part by NIH GM061068, the University of Nebraska Biochemistry Department and Redox Biology Center, the Layman Foundation, and the Nebraska Agricultural Research Division, Journal Series no. 14693. This publication was also made possible by NIH grant number P20 RR-017675-02 from the National Center for Research Resources.
The contents of this paper are solely the responsibility of the authors and do not necessarily represent the official views of the NIH.
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