An Ortholog of OxyR in Legionella pneumophila Is Expressed Postexponentially and Negatively Regulates the Alkyl Hydroperoxide Reductase (ahpC2D) Operon

Abstract

Legionella pneumophila expresses two peroxide-scavenging alkyl hydroperoxide reductase systems (AhpC1 and AhpC2D) that are expressed differentially during the bacterial growth cycle. Functional loss of the postexponentially expressed AhpC1 system is compensated for by increased expression of the exponentially expressed AhpC2D system. In this study, we used an acrylamide capture of DNA-bound complexes (ACDC) technique and mass spectrometry to identify proteins that bind to the promoter region of the ahpC2D operon. The major protein captured was an ortholog of OxyR (OxyR_Lp). Genetic studies indicated that oxyR_Lp was an essential gene expressed postexponentially and only partially complemented an Escherichia coli oxyR mutant (GS077). Gel shift assays confirmed specific binding of OxyR_Lp to ahpC2D promoter sequences, but not to promoters of ahpC1 or oxyR_Lp; however, OxyR_Lp weakly bound to E. coli OxyR-regulated promoters (katG, oxyR, and ahpCF). DNase I protection studies showed that the OxyR_Lp binding motif spanned the promoter and transcriptional start sequences of ahpC2 and that the protected region was unchanged by treatments with reducing agents or hydrogen peroxide (H₂O₂). Moreover, the OxyR_Lp (pBADLpoxyR)-mediated repression of an ahpC2-gfp reporter construct in E. coli GS077 (the oxyR mutant) was not reversed by H₂O₂ challenge. Alignments with other OxyR proteins revealed several amino acid substitutions predicted to ablate thiol oxidation or conformational changes required for activation. We suggest these mutations have locked OxyR_Lp in an active DNA-binding conformation, which has permitted a divergence of function from a regulator of oxidative stress to a cell cycle regulator, perhaps controlling gene expression during postexponential differentiation.

Legionella pneumophila is found in aquatic and moist soil environments as an intracellular parasite of protozoa (12, 14, 15). Legionnaires' disease results from aerosol transmission of bacteria from the environment to susceptible humans. In natural hosts (and some mammalian cell lines), L. pneumophila displays a dimorphic lifestyle in which vegetative replicating bacilli differentiate into metabolically dormant cyst-like planktonic forms (17). Analogous to the small-cell variants of Coxiella burnetii or the elementary bodies of Chlamydia spp., these highly resilient cyst-like forms of L. pneumophila enable the organism to survive extracellularly for extended periods of time in a highly infectious state. The cyst form likely contributes to the noted resilience of L. pneumophila to the actions of oxidizing agents (chlorine), chloramines, and biocides, used to control infection of man-made aquatic systems (17). Several lines of evidence suggest that antioxidant enzymes play crucial roles during the early stages of infection of protozoa or human phagocytes (3, 6, 8, 19, 27) and may also be important for environmental persistence (25).

We previously characterized two peroxide-scavenging alkyl hydroperoxide reductase systems of L. pneumophila: AhpC1, similar to the AhpC-thioredoxin system of Helicobacter pylori (30, 49), and AhpC2D, similar to the AhpC AhpD system found in mycobacteria (30, 42). These studies demonstrated that both systems contributed to efficient scavenging of H₂O₂ and organic peroxides and that at least one system was required for viability. Moreover, these genes were inversely expressed during the growth cycle; ahpC2D induced early in exponential phase and repressed postexponentially, concomitant with induction of ahpC1. Finally, a compensatory increase in expression of ahpC2D was observed in an ahpC1 mutant, suggesting that the ahpC2D operon might be upregulated in response to changes in cellular redox status or by increases in intracellular peroxides. However, previous studies of the antioxidant defense genes of L. pneumophila (sodB, sodC, katA, katB, ahpC1, or ahpC2D) had concluded that none of these genes were activated in response to oxidative stress (4, 5, 30, 39, 43). In contrast, these studies revealed that all of these antioxidant defense genes, with the exception of sodB (constitutive), were growth stage regulated.

The emerging pattern of growth stage-regulated antioxidant defense genes in L. pneumophila together with the postexponential expression of transmission (virulence) traits is consistent with the dimorphic lifestyle of this organism (17). In support of this view, microarray analyses comparing sessile and planktonic cells of L. pneumophila under biofilm conditions revealed ahpC2 and ahpD to be the most highly induced genes in sessile cells (25). Related studies have similarly shown that expression of the ahpC2D operon increases significantly during intracellular growth in protozoa (8). Despite these findings, little is known of the regulatory systems controlling ahpC2D expression or other antioxidant defense genes in L. pneumophila. With the exception of an OxyR homolog (described herein), typical regulators of oxidative stress defenses (such as SoxRS, OhrR, and PerR) are absent in the L. pneumophila genome. Growth phase-dependent regulation of genes associated with virulence and transmission traits has been attributed in part to global regulators, such as the stationary-phase sigma factor (RpoS) (2, 20), the carbon storage regulator (CsrA) (13, 33), and the LetAS, CpxRA, and PmrBA two-component regulatory systems (13, 16, 22, 32, 52); however, the predicted hierarchal regulatory cascade is incomplete, and none of these regulators has been linked to oxidative stress defenses.

Here we report the use of an acrylamide capture of DNA-bound complexes (ACDC) technique (35) and tandem mass spectrometry of ahpC2D promoter-bound proteins to identify a previously uncharacterized putative regulator of oxidative stress in L. pneumophila, designated OxyR_Lp. These studies will show that while oxyR_Lp was able to partly complement an oxyR mutant of Escherichia coli and purified OxyR_Lp exhibited weak binding to OxyR-regulated promoters of E. coli by gel shift experiments, the protein was no longer redox active and its repression of ahpC2D could not be reversed by H₂O₂. Finally, the apparent essentiality of oxyR_Lp together with its postexponential expression, concomitant with repression of ahpC2D, suggests regulatory roles that are unrelated to oxidative stress.

MATERIALS AND METHODS

Bacterial strains and growth conditions.

Bacterial strains and plasmids used in this study are listed in Table 1 and oligonucleotides in Table 2. L. pneumophila strains were grown aerobically at 37°C on buffered charcoal yeast extract agar or in buffered yeast extract (BYE) broth, supplemented with α-ketoglutaric acid (1 mg/ml), ferric pyrophosphate (250 μg/ml), l-cysteine (40 μg/ml), thymidine (100 μg/ml), and antibiotics when required. E. coli strains DH5α, MG1655, and GS077 were grown at 37°C on Luria-Bertani (LB) agar or in LB broth supplemented with the appropriate antibiotics. For some studies, E. coli GS077 was grown at 37°C under anaerobic conditions using the BD GasPak EZ (Becton Dickinson, Oakville, Ontario) and AnaeroGen anaerobic atmosphere generation system (Oxoid, Ltd., Nepean, Ontario). Antibiotics (Sigma-Aldrich Canada Ltd., Oakville, Ontario) were added to media at the following concentrations, when appropriate: streptomycin (100 μg/ml), kanamycin (40 μg/ml), chloramphenicol (20 μg/ml), and ampicillin (100 μg/ml). All strains were stored at −85°C in nutrient broth containing 10% dimethyl sulfoxide.

TABLE 1.

Strains, genotypes, and plasmids used in this study

Bacterial strain, genotype, or plasmid	Relevant property(ies)	Source or reference
E. coli
DH5α	F− φ80dlacZΔM15 Δ(lacZYA-argF)U169 endA1 recA1 hsdR17 deoR thi-1 supE44 gyrA96 relA1	Clontech
BL21 Codon Plus	F−ompT hsdS (rB− mB−) dcm+ Tetrgalλ (DE3) endA Hte [argU ileY leuW Camr]	Stratagene
MG1655	K-12, F− λ−	ATCC 47076
GS077	MG1655-derived oxyR::kan	51
L. pneumophila
Lp02	Philadelphia-1 derivative, rpsL hsdR thyA (Smr) mutant	7
ahpC1::kan	Lp02 ahpC1::kan mutant	30
ahpC2D::kan	Lp02 ahpC2D::kan mutant	30
rpoS::kan	Lp02 strain MB 379::kan	2
letA::kan	Lp02 strain MB 414::kan	22
himA::gm himB::kan	Lp02 integration host factor double mutant (lacking both α and β subunits); Kanr Gmr	This laboratory
Plasmids
pET29b	IPTG-inducible T7 expression vector	Novagen
petLpoxyR	pET29b-derived overexpression vector for His6-OxyRLp	This study
pBH6119	RSF1010 ori, promoterless gfpmut3 tdΔi (Ampr)	21
pC2gfp	PahpC2 region (−199 to +27 bp upstream of ahpC2) cloned upstream of gfpmut3 of pBH6119	30
pC2(−179/+27)gfp	PahpC2 region (−179 to +27 bp upstream of ahpC2) cloned upstream of gfpmut3 of pBH6119	This study
pC2(−156/+27)gfp	PahpC2 region (−156 to +27 bp upstream of ahpC2) cloned upstream of gfpmut3 of pBH6119	This study
pC2(−132/+27)gfp	PahpC2 region (−132 to +27 bp upstream of ahpC2) cloned upstream of gfpmut3 of pBH6119	This study
pC2(−109/+27)gfp	PahpC2 region (−109 to +27 bp upstream of ahpC2) cloned upstream of gfpmut3 of pBH6119	This study
pC2(−91/+27)gfp	PahpC2 region (−91 to +27 bp upstream of ahpC2) cloned upstream of gfpmut3 of pBH6119	This study
pC2(−66/+27)gfp	PahpC2 region (−66 to +27 bp upstream of ahpC2) cloned upstream of gfpmut3 of pBH6119	This study
poxyRgfp	oxyR promoter region cloned upstream of gfpmut3 of pBH6119	This study
pBAD22	Arabinose-inducible expression vector (Ampr)	18
pBADEcoxyR	pBAD22-derived expression vector for E. coli oxyR	This study
pBADLpoxyR	pBAD22-derived expression vector for L. pneumophila oxyR	This study

TABLE 2.

Oligonucleotide primers used in this study

Primer	Sequence (5′-3′)
Ac-FC2PROM	GGATCCACCAATATTCCTCCACAGGC
RC2PROM	GAATTCCTGGAAATTTGTTGCCTACCG
FECOXYRPBAD	GGCGAATTCATTATGAATCGTGATCTTGAG
RECOXYRPBAD	GGCAAGCTTAAACCGCCTGTTTTAAAACTT
FLPOXYRPBAD	GGCGAATTCATTATGAATTTAAGAGATTTACATTAT
RLPOXYRPBAD	GGCAAGCTTCTATGATAATTTGGATTGAACATT
FBLA	TCCGTGTCGCCCTTATTCC
RBLA	AACTACGATACGGGAGGGC
FLPOXYRPET	CATATGAATTTAAGAGATTTACATTAT
RLPOXYRPET	CTCGAGTGCTAATTTGGATTGAACATTTT
FC1PROM	GAATTCGTGCCATTACTGAGCGATTC
RC1PROM	GGATCCGAGTGCCTTTCATGTAGAGC
FLPOXYRPROM	GAATTCGATGGCACAAAGAGTTGCA
RLPOXYRPROM	CTGGGATCCTGCTTTACATCTGCCAGG
FECAHPCPROM	GGCACTGAAGATACCAAAGG
RECAHPCPROM	TCGATGAGATGTAAGGTAACC
FECKATGPROM	CGAAATGAGGGCGGGAAAAT
RECKATGPROM	CGATACACAGCGTTAGAGAG
FECOXYRPROM	GGTGCCGCTCCGTTTCTG
RECOXYRPROM	GCTGGCTAACGTGGCAGG
FC2PROM179	GGCGAATTCCCTTCTTTAAAAGTAGTTTATAAAA
FC2PROM156	GGCGAATTCAAACATAAAAGCCAAATATTTTGC
FC2PROM132	GGCGAATTCAAAATGGATTAATAATTCTTATATTG
FC2PROM109	GGCGAATTCTTGATTGACTTTATTTATCGTCTT
FC2PROM91	GGCGAATTCCGTCTTTATAAAAACAATTGATTTT
FC2PROM66	GGCGAATTCATTTATTAAATGATTTCACCTAAATT
RGFP	ACCATAACCGAAAGTAGTGAC

Standard molecular techniques.

Preparation of DNA, restriction enzyme digestions, and other molecular techniques have been described elsewhere (40). Plasmid isolation was performed with the QIAprep spin miniprep kit (Qiagen Inc., Mississauga, Ontario) or using a standard alkaline lysis method (40). Purification of DNA fragments from agarose gels for subcloning was carried out with a QIAquick gel purification kit (Qiagen). PCR was performed under conditions described by LeBlanc et al. (30) using oligonucleotides synthesized by Integrated DNA Technologies, Inc. (Coralville, IA). DNA and protein sequencing was carried out by the DalGen Microbial Genetics Center (Halifax, Nova Scotia, Canada). Protein concentrations were estimated using a Bio-Rad protein assay (Bio-Rad, Hercules, CA).

ACDC.

To identify transcription factors that bind to the promoter region of ahpC2D (P_ahpC2), an ACDC was performed according to a modified version of the methods described by Nelson et al. (35). Briefly, PCRs were performed using forward primers containing 5′ Acrydite (Ac) moiety, Ac-FC2PROM, and unmodified reverse primer RC2PROM. Amplicons were purified using a MinElute gel extraction kit (Qiagen). Protein lysates were obtained as follows: bacteria were harvested (4,800 × g, 6 min, 4°C) from a 500-ml, 24-h culture of L. pneumophila. The pellet was washed with cold 50 mM Tris-HCl, pH 7.5, and resuspended in buffer consisting of 50 mM Tris-HCl, pH 7.5; 1 mM EDTA, pH 8.0; 50 mM NaCl; 10% glycerol with 0.1 mM phenylmethylsulfonyl fluoride (PMSF); and 1 mM dithiothreitol (DTT). Cells were lysed using a French press and subjected to ultracentrifugation at 100,000 × g. Protein lysates were aliquoted and stored at −85°C. Before use, protein lysates were diluted in a buffer consisting of 100 mM Tris-HCl, pH 7.5, and 10 mM MgCl₂ and treated for 15 min at room temperature with 100 mM of acetyl phosphate (phosphorylate DNA-binding proteins). Binding reactions for the ACDC assay were performed in a 25-μl volume and consisted of 0 (no DNA for control), 2, 4 or 8 μg of Ac-DNA, 1 μg of protein lysate, 7.5 μl buffer D (20 mM HEPES, pH 7.8; 100 mM KCl; 10% glycerol; and 1 mM DTT), 1 μg poly(dI-dC), and 3.5 μl of 30% acrylamide. Reactions were incubated for 15 min at room temperature before addition of 1.25 μl of 5% ammonium persulfate and 1.25 μl of diluted N,N,N′,N′′-tetramethylethylenediamine (1:30 in buffer D). The reactions were immediately loaded into the wells of a precast nondenaturing 5% polyacrylamide gel (0.5× Tris-borate-EDTA [TBE], Bio-Rad Laboratories). After polymerization (approximately 5 min), electrophoresis was performed in 0.5× TBE for 1 h at 125 V to remove nonspecific proteins. Each well was excised from the gel, transferred to a microtube containing 100 μl of 10% sodium dodecyl sulfate (SDS) sample buffer (NEB) with 5% β-mercaptoethanol and heated for 3 min at 94°C to remove captured proteins. After a brief centrifugation, 50-μl aliquots were loaded onto a precast 4 to 15% Tris-HCl polyacrylamide gel (Bio-Rad Laboratories) and subjected to SDS-polyacrylamide gel electrophoresis (SDS-PAGE). After silver staining, proteins distinct from the no-DNA controls were excised and submitted for mass spectrometry analysis (DalGen Microbial Genomics Center).

Cloning and overexpression of OxyR_Lp in E. coli GS077.

For complementation of the E. coli strain GS077 (oxyR::kan) (generously donated by Gisela Storz, National Institutes of Health, Bethesda, MD), E. coli strain K-12 oxyR (oxyR_Ec) or oxyR_Lp was expressed using the arabinose-inducible expression systems of pBAD22 (18). Amplicons were generated from PCRs using primer pairs FECOXYRPBAD and RECOXYRPBAD for oxyR_Ec and FLPOXYRPBAD and RLPOXYRPBAD for oxyR_Lp, digested with EcoRI and HindIII and subcloned into pBAD22. The resulting constructs (pBADEcoxyR and pBADLpoxyR) were transformed into E. coli DH5α and verified by PCR and DNA sequencing. Plasmids were extracted and transformed into electrocompetent E. coli GS077. Ampicillin-resistant GS077 transformants obtained anaerobically were confirmed by PCR analysis and termed GS077 pBADEcoxyR and GS077 pBADLpoxyR. The pBAD22 empty vector was transformed into wild-type MG1655 and GS077 strains, verified by PCR analysis using primers for the ampicillin resistance cassette FBLA and RBLA, and termed MG1655 pBAD22 and GS077 pBAD22, respectively.

Disk diffusion assays.

Strains were grown under anaerobic conditions to mid-exponential phase (optical density at 620 nm [OD₆₂₀] of 0.4), and expression of oxyR_Ec or oxyR_Lp from respective pBAD22-derived plasmids was induced with various concentrations of arabinose (0, 0.02, and 0.2%) or repressed using 0.2% glucose. Samples were analyzed by SDS-PAGE to confirm the expression of OxyR_Ec and OxyR_Lp. Cells corresponding to 0.1 OD₆₂₀ were inoculated in 4 ml of prewarmed LB 0.75% top agar, mixed by inversion, and poured onto LB media with proper antibiotics and in the presence of arabinose or glucose. Once solidified, 10 μl of 30% H₂O₂ was placed on sterile one-fourth-inch antibiotic disks in the center of the plate. Following overnight incubation at 37°C under anaerobic conditions, the diameters of the zones of inhibition were measured and reported for three independent experiments.

Overproduction and purification of OxyR_Lp.

The primer pair FLPOXYRPET and RLPOXYRPET was used to generate an amplicon containing NdeI and XhoI restriction sites, respectively. Following digestion, the NdeI-XhoI fragment was subcloned into the corresponding sites of pET29b, generating pETLpoxyR. The in-frame fusion of L. pneumophila oxyR with the C-terminal hexameric histidine (His₆) tag was confirmed by DNA sequencing. For overexpression, an overnight culture of E. coli BL21(DE3) Codon Plus harboring pETLpoxyR was inoculated 1% (vol/vol) into 200 ml of fresh LB broth and grown to an OD₆₂₀ of 0.4. Isopropyl-β-d-thiogalactopyranoside was added to a final concentration of 1 mM, and incubation was continued for another 60 min at 37°C. Bacteria were then harvested by centrifugation (5,800 × g for 6 min), resuspended (5 ml/g [wet weight]) in suspension buffer (20 mM Tris-HCl, pH 7.9; 500 mM NaCl; 5 mM imidazole; and 1 mM PMSF), and lysed on ice by sonication (six bursts of 10 s at 200 W). Following clarification by centrifugation (10,000 × g for 15 min, 4°C), the His₆-OxyR_Lp was purified by nickel interaction chromatography as previously described (41). Samples taken from each step of purification were analyzed by SDS-PAGE, and the purified protein fractions were pooled, aliquoted, and stored at −85°C. A single band was excised and submitted for mass spectrometry analysis. Purified OxyR_Lp protein used in this study is considered to be wholly active.

EMSA.

For electrophoretic mobility shift assay (EMSA), amplicons spanning the promoter regions of ahpC1, ahpC2, and oxyR (P_ahpC1, P_ahpC2, and P_oxyR) were generated by PCR amplification using primers sets FC1PROM and RC1PROM, FC2PROM and RC2PROM, and FLPOXYRPROM and RLPOXYRPROM, respectively. For binding assays to the E. coli ahpCF, katG, and oxyR_Ec promoters, primer pairs FECAHPCPROM and RECAHPCPROM, FECKATGPROM and RECKATGPROM, and FECOXYRPROM and RECOXYRPROM were used to generate respective amplicons. Fifty nanograms of the DNA probes was mixed with various amounts of His₆-OxyR_Lp (0, 3.9, 7.8, 15.6, 31.3, 62.5, 125, 250, and 500 ng) in a 20-μl reaction mixture containing 20 mM Tris-HCl, pH 8.0; 5 mM MgCl; 20 mM KCl; 50 μg bovine serum albumin; 20% glycerol; 1 mM PMSF; and 0 or 200 mM DTT (Promega, Madison, WI) for oxidizing or reducing conditions, respectively. After 30 min of incubation at room temperature, protein-DNA complexes were separated by electrophoresis for approximately 1 h at 125 V on a 5% nondenaturing polyacrylamide gel in 0.5× TBE buffer. Gels were then soaked in 10,000-fold diluted Sybr green I nucleic acid stain (Invitrogen, La Jolla, CA) and washed twice in distilled water, and DNA was visualized using a Typhoon 9410 system (GE Healthcare) under blue light excitation at 488 nm and using a 520-nm emission filter. Densitometry was performed using ImageQuant version 5.2 (Molecular Dynamics, Sunnyvale, CA).

Mapping of P_ahpC2.

An EMSA was performed as described above using 500 ng of His₆-OxyR_Lp and 50 ng of variously sized 5′-end deletion DNA fragments generated by PCR using a common reverse primer, RC2PROM, with different forward primers: FC2PROM, FC2PROM179, FC2PROM156, FC2PROM132, FC2PROM109, FC2PROM91, and FC2PROM66. All promoter fragments were also subcloned into pBH6119 and electroporated into L. pneumophila Lp02 and the ahpC1::kan mutant (30) to evaluate promoter activity using the green fluorescence protein (GFP) reporter assay.

DNase I footprinting.

DNase I protection experiments were performed using a 226-bp DNA fragment of P_ahpC2 generated from PCRs using primer pair FC2PROM and RC2PROM (containing EcoRI and BamHI restriction sites, respectively). The digested DNA (EcoRI for the top strand and BamHI for the bottom strand) was then treated with calf intestinal phosphatase (New England BioLabs, Ipswich, MA), and reactions were purified using a QIAquick gel extraction kit (Qiagen). End-labeled DNA was prepared using [γ-³²P]dATP and Ready-To-Go T4 polynucleotide kinase (GE Healthcare) as described by the manufacturer and purified with a QIAquick nucleotide removal kit (Qiagen). Binding reactions were performed by incubating approximately 20,000 cpm of labeled probe with 10 μg of OxyR_Lp protein in an assay buffer consisting of 20 mM Tris-HCl, pH 8.0; 10 mM MgCl₂; 100 mM KCl; 1 mM CaCl₂; 10% glycerol; 0.05 mg/ml bovine serum albumin; 4 μg/ml poly(dI-dC); and H₂O₂ (0 to 50 mM) for oxidizing or 200 mM DTT for reducing conditions. In cases where peroxidation abolished DNA binding, replicates were treated with 200 mM DTT to restore DNA binding activity. After 20 min of incubation, 0.5 to 1 U of DNase I (Sigma) was added. The reaction was continued for 90 s before 100 μl of a stop solution (200 mM EDTA, pH 8.0; 5 M ammonium acetate; and 100 μg of salmon sperm DNA) was added. DNA was ethanol precipitated following phenol-chloroform extraction. Dry pellets were mixed with a formamide loading dye and loaded onto a 6% sequencing gel. The DNA ladder was performed by cycle sequencing using an α-³⁵S-dATP label according to the SequiTherm Excel II DNA sequencing kit (Epicenter Technologies).

Primer extension.

Total RNA was obtained from exponentially grown L. pneumophila by an automated Maxwell-16 protocol (Promega), and primer extension reactions were performed as follows. Briefly, 2 pmol of [γ-³²P]ATP end-labeled RC2PROM primer was added to 25 μg total RNA and allowed to anneal by dropping the temperature from 80°C to 41°C over a period of 30 min. cDNA extension products were generated at 41°C for 1 h in a reaction mixture containing 1 μl of 200 U/μl SuperScript reverse transcriptase (Invitrogen) and subsequently treated with RNase A (Sigma) to eliminate residual RNA. Samples were loaded onto a 6% polyacrylamide urea sequencing gel along with the corresponding DNA sequence ladder obtained by cycle sequencing using an α-³⁵S-dATP label with the appropriate primer in accordance with the SequiTherm Excel II DNA sequencing kit protocol.

GFP reporter assay.

A GFP-transcriptional fusion of P_oxyR was constructed using primer pair FLPOXYRPROM and RLPOXYRPROM and cloned into the EcoRI and BamHI sites of pBH6119 (21). Plasmid constructs were first transformed into E. coli DH5α strains, and correct orientation was determined by PCR using the respective combination of FLPOXYRPROM and RGFP primers. The pBH6119 plasmid or its derivatives were electroporated into wild-type, ahpC1, ahpC2D, rpoS, letA, or himAB mutants of L. pneumophila. Transformants capable of growth without thymidine were verified for respective plasmids by PCR analysis using primer pairs FLPOXYRPROM and RLPOXYRPROM or FBLA and RBLA for the empty vector controls. Fluorometric detection of samples taken from BYE-grown bacteria was performed as described previously (30). Values are expressed as relative fluorescence units (RFU) per OD₆₂₀ and represent triplicate values obtained from three independent experiments.

OxyR_Lp regulation of ahpC2-gfp in E. coli GS077.

DNA fragments containing gfp (no promoter) or P_ahpC2gfp were excised with EcoRI and PstI from plasmids pBH6119 and pC2gfp, respectively, and were subcloned into pMMB206. The resulting constructs, pgfp and pahpC2gfp, respectively, were electroporated into either GS077 pBAD22 or GS077 pBADLpoxyR. Ampicillin- and chloramphenicol-resistant transformants were confirmed by PCR analysis and subjected to the GFP reporter assay (30). Overnight cultures, grown at 37°C in LB broth under anaerobic conditions, were inoculated 1% into fresh media until an OD₆₂₀ of 0.4 was obtained. Then, glucose or arabinose was added to a final concentration of 0.02%, and cultures were incubated for an additional hour. In some cases, strains were then challenged for 30 min with 100 μM of H₂O₂. Fluorometric detection of samples taken from LB-grown bacteria was performed as described previously (30). Values are expressed as RFU per OD₆₂₀ and represent triplicate values obtained from three independent experiments.

RESULTS

Acrylamide capture of ahpC2D DNA-bound complexes.

We employed the ACDC technique (Fig. 1) to screen for transcription factors that bound the promoter region of ahpC2D. The principle of this method relies on capture of DNA-binding proteins from L. pneumophila lysates by interaction with ahpC2D promoter DNA sequences immobilized in the polyacrylamide matrix by covalent linkage to its 5′ Ac moiety. Proteins that do not bind the Ac-DNA can be removed by standard nondenaturing electrophoresis. Following SDS-PAGE, silver staining, and mass spectrometry analysis of proteins captured in the wells of the native acrylamide gel, proteins able to bind target DNA sequences can be identified. Using increasing concentrations of Ac-labeled ahpC2D promoter DNA (Ac-P_ahpC2) in the ACDC assay (Fig. 1 inset), a distinct protein was captured by the immobilized Ac-DNA, whereas no bands could be seen in the absence of Ac-DNA. Peptides identified by tandem mass spectroscopy analysis corresponded to 181 of 296 amino acids (61% sequence coverage) of a previously uncharacterized protein annotated as the hydrogen peroxide-inducible gene activator, OxyR, of L. pneumophila Philadelphia-1. This protein, designated OxyR_Lp, shared 45 to 50% identity and 63 to 69% similarity to well-characterized OxyR regulators of Xanthomonas campestris, E. coli, Salmonella enterica serovar Typhimurium, Haemophilus influenzae, and C. burnetii by BLAST analysis.

FIG. 1.

Acrylamide capture of proteins interacting with P_ahpC2. The ACDC assay was performed by adding 1 μg of protein (cell extract) prepared from BYE-grown L. pneumophila to various amounts of Ac-DNA (0, 2, 4, and 8 μg). Nonspecific proteins were removed by nondenaturing electrophoresis on a 0.5× TBE-5% polyacrylamide gel. Captured proteins (inset) were excised from wells, heated in SDS sample buffer containing β-mercaptoethanol, and following SDS-PAGE, visualized by silver staining. dsDNA, double-stranded DNA; APS, ammonium persulfate; TEMED, N,N,N′,N′′-tetramethylethylenediamine.

The identification of OxyR_Lp as a putative regulator of ahpC2D was not unexpected, since OxyR mediates oxidative stress induction of alkyl hydroperoxide reductase and catalase genes in many organisms (1, 26, 31, 36). Moreover, in those organisms expressing the ahpC2D operon, including close relative C. burnetii, oxyR is usually located immediately upstream and shares promoter and regulatory sequences. However, the oxyR_Lp gene was not located upstream of ahpC2D, but rather shares a divergent promoter region with a gene annotated as a major facilitator transporter, and downstream is a gene of unknown function oriented in the opposite direction. Our attempts to knock out oxyR_Lp by allelic replacement with a drug resistance cassette (kanamycin) by methods previously used to obtain ahpC1 and ahpC2D mutants (30) proved unsuccessful. Since catalase supplementation to media enabled the recovery of oxyR mutants in Pseudomonas aeruginosa (23), buffered charcoal yeast extract agar was supplemented with catalase (10,000 units); however, no oxyR_Lp mutants were recovered. The apparent essentiality of oxyR_Lp was unexpected, since the genomic organization was uncomplicated by overlapping regulatory elements or downstream polarity effects. Possible functional divergence was considered, since previous studies had indicated that genes typically regulated in response to oxidative stress in other bacteria (ahpC1, ahpC2D, katA, katB, sodB, and sodC) were not similarly regulated in L. pneumophila (4, 5, 30, 38, 39, 43). Rather, most of these genes exhibited growth stage-dependent expression (4, 5, 30, 43).

Complementation of E. coli oxyR mutant.

Functional complementation was assessed by cloning (pBAD22) and expressing the oxyR_Lp gene in E. coli oxyR mutant strain GS077. Studies were conducted under anaerobic conditions because this strain is highly susceptible to peroxide killing (compared with wild-type strain MG1655) due to a loss of OxyR regulatory functions. Under inducing conditions (arabinose) or repressing conditions (glucose), the presence of pBAD22 did not alter peroxide susceptibility (Table 3). Complementation of GS077 with pBADEcoxyR fully restored peroxide resistance to wild-type E. coli MG1655 levels under inducing conditions, but not under repressing conditions. In contrast, overexpression of pBADLpoxyR partially rescued the peroxide sensitivity of E. coli GS077, suggesting that OxyR_Lp recognizes OxyR-regulated promoters. The noted partial complementation by OxyR_Lp might be due to DNA sequence differences in OxyR binding motifs between these organisms.

TABLE 3.

Zones of inhibition (mm) in a disk diffusion assay using 10 μl of 3% H₂O₂

Straina	Zones of inhibition (mm) with:
	Glucose (%)		Arabinose (%)b
	0	0.2	0.02	0.2
K-12†	21 ± 1	23 ± 1	21 ± 1	21 ± 1
K-12 pBAD22‡	21 ± 1	22 ± 1	22 ± 1	22 ± 1
GS077§	38 ± 2	40 ± 1	38 ± 1	37 ± 2
GS077 pBAD22‡	39 ± 1	41 ± 1	38 ± 1	38 ± 1
GS077 pBADEcoxyR**	37 ± 2	40 ± 1	19* ± 1	18* ± 1
GS077 pBADLpoxyR**	39 ± 1	39 ± 1	29* ± 2	29* ± 2

^aTriplicate experiments were performed under anaerobic conditions on Mueller-Hinton agar without antibiotics (†) or supplemented with 100 μg/ml ampicillin (‡), with 40 μg/ml kanamycin (§), or with 100 μg/ml ampicillin and 40 μg/ml kanamycin (**).

^b*, significance (P < 0.01) was determined in relation to strain GS077.

Interaction of OxyR_Lp with the promoter region of ahpC2.

To both confirm ACDC results and extend functional studies, Ni²⁺-nitrilotriacetic acid-purified OxyR_Lp (overexpression from pETLpoxyR) was assessed by EMSA for DNA-binding specificity to the promoter regions (P_ahpC2, P_ahpC1, and P_oxyR) of these genes. As seen in Fig. 2A, OxyR_Lp bound to a 226-bp DNA fragment containing P_ahpC2 (−199 to +27 relative to the ahpC2 coding sequence). This is evident in the sample containing 500 ng of His₆-OxyR_Lp, where 84% of the DNA probe was in the bound state. No differences in binding were observed under oxidizing (no added reducing agent) or reducing conditions (200 mM DTT) (see Fig. 2B). Similar experiments were also carried out with P_ahpC1 and P_oxyR; however, no DNA binding was observed (data not shown). EMSAs were also performed with amplicons containing promoter regions of known members of the E. coli OxyR regulon (ahpCF, katG, and oxyR) under oxidizing conditions. As seen in Fig. 2C, OxyR_Lp weakly bound these promoters and only at the highest concentrations of protein used (500 ng). The EMSA results indicate that the partial complementation of an E. coli oxyR mutant by oxyR_Lp might be due to a combination of OxyR_Lp overexpression and partial activation of OxyR-regulated promoters in GS077.

FIG. 2.

Interaction between OxyR_Lp and P_ahpC2. (A) Fifty ng of a 226-bp P_ahpC2 fragment spanning the region −199 to +27 (relative to the L. pneumophila ahpC2 initiation codon) was incubated in the presence of various amounts of His₆-OxyR_Lp. Lanes: 1, low-mass ladder (Invitrogen); 2, DNA without protein; 3 to 10, DNA in the presence of 3.9, 7.8, 15.6, 31.3, 62.5, 125, 250, or 500 ng of His₆-OxyR_Lp, respectively. Arrows indicate bands representing the DNA probe (P) and mobility shift (S). Densitometry analysis reported as a percentage of DNA probe (P [%]) or shift (S [%]) relative to the total intensity of each lane. (B) Mobility shift assay performed in the presence of 25 ng P_ahpC2 with OxyR_Lp under reducing (200 mM DTT, OxyR_Lp-red) or oxidizing (no DTT, OxyR_Lp-ox) conditions. (C) Mobility shift assay performed in the presence of 50 ng each of E. coli P_ahpCF, P_katG, and P_oxyR in the presence (+) or absence (−) of 500 ng His₆-OxyR_Lp under nonreducing conditions.

Mapping of the OxyR binding site within P_ahpC2.

To identify upstream DNA sequences necessary for ahpC2 expression, PCR-generated 5′-end deletion fragments of P_ahpC2 were cloned upstream of gfpmut3, encoding GFP. The promoter fragments shared a common 3′ end at +27 (relative to the ahpC2 initiation codon) and a variable 5′ end ranging from −199 to −66 (Fig. 3A). The ability of P_ahpC2 fragments to express GFP was assessed in the ahpC1 mutant (Fig. 3B), since ahpC2 levels are increased in this mutant (30). The 5′-end deletions of P_ahpC2 from base pairs −179 to −109 showed transcriptional activity similar to that of the previously studied full-length fragment (−199 to +27) (30). However, P_ahpC2 deletions of −91 or −66 no longer expressed GFP. It should be noted that similar results were obtained in the wild-type strain, albeit at a lower expression level (data not shown). In a parallel series of experiments, mobility shift assays were performed with various P_ahpC2 fragments (Fig. 3C). Deletion of nucleotides −199 to −109 had no effect on OxyR_Lp binding (Fig. 3C, lanes 2 to 5). However, further deletion of nucleotides −91 to −66 (which had ablated transcription) did not completely abolish DNA binding, suggesting that OxyR_Lp binding motif(s) remained. An alignment of DNA sequences for the −105 to −55 region of P_ahpC2 for the three genome sequences for L. pneumophila together with sequences from OxyR-regulated genes from E. coli and other species reveals some conservation with the E. coli OxyR consensus (Fig. 3D). Taken together, these findings suggest that the OxyR_Lp-protected region contains an OxyR motif that might include promoter sequences.

FIG. 3.

Promoter deletion analysis of the OxyR_Lp binding site within P_ahpC2. (A) Schematic representation of the 5′-end deletions of P_ahpC2 cloned upstream of promoter-less gfp. (B) Fluorescence detected from ahpC1::kan mutants harboring plasmids and P_ahpC2 promoter DNA-GFP constructs. (C) Mobility shift assay performed with the 5′-end deletions of P_ahpC2. Each lane contained 50 ng of the DNA fragments incubated with (right panel) or without (left panel) 500 ng of His₆-OxyR_Lp. For both gels, lanes are as follows (symbols as shown in panel B): 1, low-mass ladder (Invitrogen); 2, P_ahpC2 fragment −199 to +27 (▪); 3, P_ahpC2 fragment −179 to +27 (⋄); 4, P_ahpC2 fragment −156 to +27 (▴); 5, P_ahpC2 fragment −132 to +27 (*); 6, P_ahpC2 fragment −109 to +27 (○); 7, P_ahpC2 fragment −91 to +27 (•); 8, P_ahpC2 fragment −66 to +27 (▵); and 9, no-DNA control (⧫). DNA fragments are annotated relative to the ahpC2 initiation codon. (D) DNA alignments of the ahpC2 upstream region (−115 to −55) from L. pneumophila Philadelphia-1, L. pneumophila Paris, and L. pneumophila Lens; ahpC from Xanthomonas campestris (Xc) and P. aeruginosa (Pa); and ahpC, katG, oxyR, and oxyS from E. coli. The consensus sequence from E. coli is provided, and the matching motifs are underlined.

DNase I footprinting and primer extension.

To further delineate the specific DNA binding sequence of P_ahpC2 recognized by OxyR_Lp, the untreated and reduced (200 mM DTT) forms of OxyR (OxyR_Lp-ox and OxyR_Lp-red, respectively) were used in a footprinting assay. OxyR_Lp-red and OxyR_Lp-ox showed similar DNA footprints (Fig. 4 A and B) and confirmed that the protected region included the −91 and −66 regions. Primer extension analysis (Fig. 4C) revealed that the transcription start site as well as the −10 hexamer was located within the OxyR_Lp-protected region (Fig. 4D). The location of the promoter region and transcription start site within the OxyR_Lp-protected region is unusual, since in E. coli, reduced OxyR binds upstream of the promoter region of OxyR-regulated genes and represses transcription by overlapping the −35 hexamer (9, 31, 34, 36, 45, 46). In the case of P_ahpC2, the −35 hexamer is outside of the OxyR_Lp-protected region, suggesting a different regulatory mechanism from that reported for OxyR_Ec (46, 47). However, if OxyR_Lp was not fully oxidized during the purification process, as is typical for other OxyR proteins (47, 48), then perhaps further oxidation with H₂O₂ might reveal more typical redox-mediated DNA-binding differences.

FIG. 4.

OxyR_Lp binding of P_ahpC2 and ahpC2 transcript start site. Shaded bars indicate OxyR_Lp DNA footprints for bottom strand (end labeled at BamHI) (A) and top strand (end labeled at EcoRI) (B). OxyR_Lp binding to P_ahpC2 occurred under the following conditions: lanes 1 and 6, no protein control reactions; lane 2, oxidizing conditions (no DTT); lane 3, 200 mM DTT; lanes 4, 1 mM H₂O₂; lane 5, treated with 1 mM H₂O₂ and then with 200 mM DTT to recover DNA-binding activity. In panels A and C, asterisk denotes ahpC2 transcript start site as indicated by alignment of primer extension product with the corresponding DNA sequence ladder. (D) Schematic of P_ahpC2. Arrows indicate FC2PROM and RC2PROM primers, open boxes denote OxyR_Lp sites as determined by DNase I protection assays shown in panels A and B, boldface indicates open reading frames of ahpC2 and lpg2351, and the ribosome-binding site (RBS) for ahpC2 is underlined. The transcription start site is indicated by an asterisk, and the −10 sequence is indicated in bold. The numbers in parentheses depict the numbering and sites of deletions shown in Fig. 3.

To test this possibility, we treated OxyR_Lp with a range of peroxide concentrations (10 μM to 50 mM) and determined that 1 mM completely abolished DNA binding (Fig. 4A, lane 4). While not shown, treatment of OxyR_Lp with 500 μM H₂O₂ reduced DNA binding by 50% but did not alter the protected area relative to untreated OxyR_Lp controls. To distinguish between reversible and irreversible oxidation of OxyR_Lp, we followed 1 mM H₂O₂ treatment with 200 mM DTT treatment to reduce the disulfide bonds, and as seen in Fig. 4A, lane 5, subsequent reduction could not restore DNA binding activity. These studies indicate that the OxyR_Lp footprint is unchanged by treatments with H₂O₂ at concentrations that should be sufficient to fully oxidize the protein.

OxyR_Lp represses ahpC2-gfp expression.

Since the OxyR_Lp footprint for ahpC2 appears to block the binding of the RNA polymerase complex, we sought to confirm that OxyR_Lp was indeed a repressor. We first introduced pahpC2gfp into E. coli GS077 (the oxyR mutant) and noted constitutive expression of GFP (Fig. 5). We then introduced pBADLpoxyR and pBAD22 vector controls into GS077 strains harboring pahpC2gfp and pgfp controls and induced each with either arabinose or glucose. As seen in Fig. 5, arabinose-dependent induction of OxyR_Lp from pBADLpoxyR repressed fluorescence from pahpC2gfp, consistent with our hypothesis that binding of OxyR_Lp to the promoter of ahpC2-gfp blocked transcription. Moreover, there was no change in ahpC2-gfp expression following challenge with 100 μM H₂O₂ (oxidative stress), further evidence that OxyR_Lp is not a redox-active regulator.

FIG. 5.

Regulation of ahpC2-gfp in E. coli GS077 by OxyR_Lp. Strains were grown in LB broth and subjected to repressing (glucose) or inducing (arabinose) conditions. Treatment with 100 μM H₂O₂ is indicated where appropriate. RFUs were normalized to 1.0 OD₆₂₀ unit. Results are the means ± standard deviations of three independent experiments.

Cell cycle expression of oxyR_Lp.

To determine the temporal expression of oxyR during in vitro growth, a GFP reporter assay was performed. Figure 6 shows that the expression of oxyR_Lp is growth cycle-dependent, increasing during mid- and postexponential phases and decreasing slightly during late-stationary phase. Similar trends were observed in peroxide-sensitive ahpC1 and ahpC2D mutants. In an attempt to identify possible regulators of oxyR expression, the poxyRgfp reporter construct was transformed into mutant strains lacking known regulators of L. pneumophila virulence, rpoS, letA, and himAB (encoding integration host factor). No differences in oxyR expression were observed in any of the mutants analyzed (Fig. 6), indicating that the postexponential expression of OxyR_Lp is not activated by RpoS or the LetA/S regulatory cascade.

FIG. 6.

Cell cycle-dependent oxyR expression in L. pneumophila. (A) GFP reporter assays and (B) bacterial growth curves for P_oxyR were performed in BYE broth, and fluorescence was determined for samples taken every 3 h, following the growth of the wild type (⧫) or the ahpC2D (•), rpoS (▵), letA (▴), and himAB (encoding integration host factor) (○) mutant strains. RFUs were normalized to 1.0 OD₆₂₀ unit. Results are the means ± standard deviations of three independent experiments.

DISCUSSION

Our laboratory has characterized two alkyl hydroperoxide reductases (AhpC1 and AhpC2D) that are expressed at different stages of bacterial growth and provide essential peroxidatic functions in L. pneumophila. Loss of AhpC1 function results in increased expression of ahpC2D, suggesting a compensatory response to cellular oxidative stress (30). Using an ACDC assay, we identified OxyR_Lp as a regulatory factor controlling expression of the ahpC2D operon. OxyR_Lp bound to ahpC2D promoter sequences but did not bind to the promoter regions of ahpC1 or itself (autoregulation). Members of the OxyR family are well-characterized, redox-active transcriptional activators of genes associated with defense against oxidative stress. In contrast, none of the antioxidant defense genes (sodB, sodC, katA, katB, or ahpC1 and ahpC2D) of L. pneumophila appear to be activated in response to oxidative stress (4, 5, 30, 39, 43). Moreover, the expression of most of these genes, including oxyR_Lp, is aligned with growth stage, increasing dramatically around mid-exponential phase and peaking in early stationary phase, consistent with the view that oxidative stress is more acute for stationary-phase bacteria (4, 39). By demonstrating that OxyR_Lp repressed transcription from ahpC2D, we have partly resolved the basis for the reciprocal gene expression noted between ahpC1 and ahpC2D. Studies are in progress to identify regulatory mechanisms associated with postexponential activation of ahpC1 and oxyR.

OxyR_Lp appears to be functionally different from other OxyR proteins that have been shown to function as redox-active repressors (47, 48). Here we present several lines of evidence to suggest that OxyR_Lp may no longer function as a global regulator of antioxidant defenses: (i) oxyR_Lp expression is growth stage dependent, (ii) OxyR_Lp function is apparently essential, (iii) DNA-binding properties of OxyR_Lp are not redox dependent, (iv) OxyR_Lp contains key amino acid substitutions that likely ablate disulfide-bond formation or conformational changes that are required for activation, and (v) H₂O₂ treatment did not increase ahpC2-gfp levels in L. pneumophila or in an E. coli oxyR mutant (GS077) expressing oxyR_Lp. However, these findings do not exclude the possibility that OxyR_Lp retains some DNA sequence recognition for OxyR-regulated promoters of E. coli, as indicated by complementation and EMSA studies. Our studies indicate that OxyR_Lp may be locked in an activated DNA-binding conformation (29), independent of further activation by peroxide but sufficient to activate the antioxidant defense system of E. coli.

In E. coli, OxyR binds DNA as a dimer of dimers under both reducing and oxidizing conditions, yet only the latter promotes expression of its target genes (47). Treatment of E. coli OxyR with millimolar amounts of DTT was shown to promote the binding of OxyR to contact sites that blocked −35 sequences of OxyR-regulated promoters, whereas peroxide oxidation of OxyR led to a conformational change in the protein that contracted the DNA footprint and permitted strong induction of these promoters (44, 46, 47). In the case of L. pneumophila, treatment of OxyR_Lp with micromolar concentrations of H₂O₂ did not alter the footprint, which suggested either that OxyR_Lp was fully oxidized during protein purification or that DNA binding was independent of oxidation state. While millimolar concentrations of H₂O₂ indeed abolished DNA binding, the result is not considered physiologically relevant since biological activity was not recoverable by treatments with 200 mM DTT. In this regard, functional studies have shown that the redox-active thiols (C199 and C208) of OxyR_Ec spontaneously oxidize to form disulfide bonds (redox potential of −185 mV) during protein purification (1, 50). In some organisms, such as Deinococcus radiodurans, oxidation of a single (peroxidatic) cysteine residue of OxyR is sufficient to promote activation (10). Thus, regardless of whether a single cysteine becomes oxidized or even a disulfide bond is formed, our studies suggest that the redox-sensing domain of OxyR_Lp no longer signals conformational changes that promote activation.

Members of the LysR family (which includes OxyR) use a ligand-binding conformational change mechanism to mediate DNA binding (11). Mutation-based studies have identified key amino acids required for the function of OxyR. For example, E. coli OxyR H125I and H218D mutations resulted in wild-type DNA binding but a loss of redox-mediated regulation (11). These amino acid substitutions occur naturally in the OxyR_Lp proteins from all four strains of L. pneumophila whose genomes have been sequenced (125 and 218) (Fig. 7). While the two redox-active cysteine residues involved in OxyR activation (Cys199 and Cys208) are conserved in OxyR_Lp, there is substantial amino acid sequence divergence in the activation domain that might affect redox activity. For example, a key arginine at position 220 in OxyR_Ec (Fig. 7) that is predicted to interact with catalytic cysteines and peroxide (28) is changed to threonine in OxyR_Lp proteins. Interestingly, in the OxyR proteins of close relative C. burnetii and in Francisella tularensis, Arg220 is replaced by lysine, which might suggest a more traditional function in these organisms. Further comparisons revealed amino acid sequence variation within the helix-turn-helix region that might account for differences in the DNA-binding specificity noted between OxyR_Lp and OxyR_Ec in complementation experiments and EMSAs. We suggest that the loss of the redox-sensing function of OxyR_Lp has occurred naturally through specific amino acid substitutions that have been demonstrated through empirical studies to alter disulfide bond formation and conformational changes required for activation.

FIG. 7.

Clustal W alignment of selected OxyR amino acid sequences. The alignments are of L. pneumophila Philadelphia-1 (LpnOxyR [OxyR_Lp]), Xanthomonas campestris (XanOxyR [OxyR_Xc]), E. coli (EcoliOxyR [OxyR_Ec]), and C. burnetii (CbOxyR [OxyR_Cb]). The DNA binding helix-turn-helix (HTH) is shaded, and relevant amino acid positions are numbered and bolded.

In contrast to other bacteria, such as F. tularensis, C. burnetii, and Streptomyces species, in which oxyR is located upstream and divergently expressed from the ahpC2D system, the oxyR_Lp gene is located elsewhere in the chromosome. It is conceivable that the present oxyR_Lp was acquired laterally, perhaps concomitantly with the deletion or loss of an ancestral upstream ortholog, perhaps a necessary event associated with the evolution of a dimorphic lifestyle.

It seems unlikely that the only function for OxyR_Lp is to repress ahpC2D during the growth transition to stationary phase and into cyst morphogenesis. While downregulation of ahpC2D is consistent with metabolic changes and a shift to the NADPH-thioredoxin AhpC1 system that occur during the transition to stationary phase, ahpC2D deletion mutants are both viable and infectious (30). Moreover, high-level ahpC2D expression, noted with the ahpC1 deletion mutant, also suggested that the postexponential presence of AhpC2D was not toxic. Thus, OxyR_Lp must provide other essential regulatory functions to L. pneumophila, since this mutant could not be obtained by methods routinely used to generate mutants (30). We used the pattern search option of the L. pneumophila Paris and Lens genome website (http://genolist.pasteur.fr/LegioList/) to screen for additional genes, using motif patterns deduced from comparisons with either the E. coli or L. pneumophila OxyR binding motif. While these motifs are relatively degenerate, binding sites were observed upstream of approximately 50 genes, including the dps homolog, which has recently been found to prevent Fenton-mediated DNA damage by sequestering iron (37). Others included promoters of genes encoding efflux pumps (that exclude redox-cycling compounds and organic solvents); metal ion transporters (which regulate rates of Fenton chemistry); components of the respiratory chain, such as cytochrome oxidases, DNA repair, or modification enzymes; known virulence factors, such as the zinc metalloproteinase (shown to inhibit the oxidative burst); and substrates of the Dot/Icm type IVB secretion system (SidG, SdhB, and SdeA). Finally, OxyR binding motifs were also found upstream of genes encoding numerous transcriptional regulators, like Fur, FleQ, FleN, and other members of the LysR family, suggesting that there could be cross talk with other regulatory pathways. Interestingly, unlike in E. coli, fur is an essential gene in L. pneumophila (24). Further studies will be required to dissect additional regulatory functions associated with essentiality.

Our studies predict the participation of additional regulatory elements in controlling the expression of ahpC2D, since repression of ahpC2D by OxyR_Lp does not fully explain the upregulation of ahpC2D in an ahpC1 mutant or the postexponential activation of ahpC1 (30). In L. pneumophila, transition from exponential phase to stationary phase/cyst differentiation is controlled in part by postexponential regulators RpoS, LetA, and HimAB (2, 13, 20, 22, 32, 33, 52). However, the oxyR_Lp-gfp gene expression studies of rpoS, letA, and himAB mutants were indistinguishable from those of the wild type. While not directly examined, carbon storage regulator CsrA, an important repressor of stationary-phase genes during exponential phase (13, 33), may not be involved in regulating oxyR_Lp, since CsrA should have repressed postexponential expression of oxyR_Lp in letA and rpoS mutants. Future studies will employ the ACDC strategy to capture putative regulatory factors associated with the control of oxyR_Lp expression.

In summary, we have shown that OxyR_Lp represses ahpC2D expression and that oxyR_Lp is induced postexponentially. Our studies further establish that OxyR_Lp no longer functions as an oxidative response regulator in L. pneumophila, which is consistent with previous observations that L. pneumophila mounts no response to oxidative stress and that these bacteria are rather resistant to oxidative damage, even in phagocytic cells (30). We propose that OxyR_Lp has been adapted from an oxidative stress response regulator to a growth stage-specific regulator of genes mediating the transition from vegetative to resilient cyst-like transmissible forms.