Discovery of Catalases in Members of the Chlamydiales Order

Brigida Rusconi, Gilbert Greub

Institute of Microbiology, University of Lausanne and University Hospital Center, Lausanne, Switzerland

Catalase is an important virulence factor for survival in macrophages and other phagocytic cells. In Chlamydiaceae, no catalasehad been described so far. With the sequencing and annotation of the full genomes of Chlamydia-related bacteria, the presenceof different catalase-encoding genes has been documented. However, their distribution in the Chlamydiales order and the func-tionality of these catalases remain unknown. Phylogeny of chlamydial catalases was inferred using MrBayes, maximum likeli-hood, and maximum parsimony algorithms, allowing the description of three clade 3 and two clade 2 catalases. Only monofunc-tional catalases were found (no catalase-peroxidase or Mn-catalase). All presented a conserved catalytic domain and tertiarystructure. Enzymatic activity of cloned chlamydial catalases was assessed by measuring hydrogen peroxide degradation. Thecatalases are enzymatically active with different efficiencies. The catalase of Parachlamydia acanthamoebae is the least efficientof all (its catalytic activity was 2 logs lower than that of Pseudomonas aeruginosa). Based on the phylogenetic analysis, we hy-pothesize that an ancestral class 2 catalase probably was present in the common ancestor of all current Chlamydiales but wasretained only in Criblamydia sequanensis and Neochlamydia hartmannellae. The catalases of class 3, present in Estrella lausan-nensis and Parachlamydia acanthamoebae, probably were acquired by lateral gene transfer from Rhizobiales, whereas for Wad-dlia chondrophila they likely originated from Legionellales or Actinomycetales. The acquisition of catalases on several occasionsin the Chlamydiales suggests the importance of this enzyme for the bacteria in their host environment.

Macrophages are often a target of intracellular bacteria (1).The bacteria can be obligate intracellular bacteria, like Rick-

ettsia spp. (2), or facultative intracellular bacteria, such as Legion-ella pneumophila (3), Brucella abortus (4), and Mycobacterium tu-berculosis (5). Among members of the Chlamydiales order,Chlamydia trachomatis (serovar K9) does not resist macrophagemicrobicidal effectors (6), whereas Waddlia chondrophila is ableto replicate very efficiently in macrophages (7, 8) and Parachla-mydia acanthamoebae is able to replicate to a lower extent, rapidlyinducing the apoptotic death of the macrophage (9–11).

The ability to grow in a professional phagocyte offers severaladvantages to the invading pathogen. First, macrophages repre-sent an interesting cell target due to their presence in almost alltissues. Moreover, infecting macrophages give the bacteria an op-portunity to hamper macrophage activation, which delays the de-velopment of an effective cytotoxic T cell response. Therefore, it iscrucial to determine which factors define the resistance of macro-phages to certain bacteria, especially when macrophages exhibiteda different permissivity to bacteria belonging to the same order.

One of the first lines of defense of macrophages is the rapiddegradation of the bacteria by acidic pH, lysosomal hydrolases,and various other microbicidal effectors, including reactive oxy-gen species (ROS) produced by the transmembranous NADPHoxidase complex (NOX2). Catalases are bacterial enzymes thatdegrade ROS (H2O2). Catalases belong to a very diverse functionalclass of proteins that can be classified in four main groups: theheme-containing monofunctional catalases, the heme-containingbifunctional catalase-peroxidases, nonheme catalases, and unclas-sified catalases (12). Mutations in certain components of theNOX2 complex cause an immune disease called chronic granulo-matous disease (CGD) (13). This genetic disease is associated withrecurrent bacterial and fungal infections. Interestingly, pathogensthat infect CGD patients are expressing a ROS-degrading enzymecatalase (14) belonging to the heme-containing mono- or bifunc-tional catalase group.

Catalase-peroxidases are thought to have been acquiredthrough lateral gene transfer from archaea (15), while monofunc-tional heme-containing catalases are believed to be very ancient.The latter can be further subdivided into three main clades.

During annotation of the genome of Waddlia chondrophila inour group, a catalase-encoding gene was identified (16). Sincenone of the members of the Chlamydiaceae family encode a cata-lase, we investigated the presence of these genes in other membersof the Chlamydiales order, including Waddliaceae, Parachlamydi-aceae, Simkaniaceae, and Criblamydiaceae families. The functionalproperties and evolutionary history of identified catalases werethen assessed.

MATERIALS AND METHODSPhylogenetic analysis. BLASTP was performed on the NCBI BLAST plat-form (17). Protein sequences used for searches were retrieved fromPubMed and Uniprot. Sequences were aligned with MUSCLE (18) inGeneious v6. Domains were determined with PROSITE (19, 20). Theconsensus sequence display was created with WebLogo (21).

Prior to phylogenetic analyses, the amino acid substitution model wasassessed with ProtTest v.3.2 and modelgenerator (22). Phylogenetic anal-ysis was done with the following models. Maximum likelihood (ML) wasperformed with the PhyML 3.0 platform using the LG substitution matrix,with invariant gamma distribution (4 categories) and 500 bootstraps (23).The maximum parsimony (MP) tree was obtained using the close-neigh-bor-interchange algorithm (24) with search level 1, in which the initial

Received 14 May 2013 Accepted 30 May 2013

Published ahead of print 31 May 2013

Address correspondence to Gilbert Greub, Gilbert.Greub@chuv.ch.

Supplemental material for this article may be found at http://dx.doi.org/10.1128/JB.00563-13.

Copyright © 2013, American Society for Microbiology. All Rights Reserved.

doi:10.1128/JB.00563-13

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trees were obtained with the random addition of sequences (10 replicates)and 500 bootstraps. Evolutionary analyses were conducted in MEGA5(25). A third tree was constructed with MrBayes using LG and 1,000,000generations with invariant gamma distribution (4 categories) and fre-quency (26). The quality of the Bayesian phylogenetic tree was assessed byAWTY (J. C. Wilgenbusch, D. L. Warren, and D. L. Swofford, 2004 [http://ceb.csit.fsu.edu/awty]). Trees were visualized with FigTree v1.3.1.

Protein modeling. Protein structures were modeled with 3DJIGSAW(28–30) and SWISS-MODEL Workspace (31–34). The quality of themodels obtained with SWISS-MODEL Workspace was assessed withQMEAN4 (35, 36). The model then was further analyzed in Deepview(32). Other clade 3 catalases from P. mirabilis (Protein Data Bank [PDB]code 1M85) (37) and E. faecalis (PDB code 1SI8) (38) were used as con-trols. Root mean square (RMS), threading energy, and conformationclashes were determined with Deepview (32). For clade 2 catalases, Esch-erichia coli HPII PDB codes 1GGE (39) and 1IPH (40) were used formodeling.

Catalase cloning and expression. Genomic DNA of W. chondrophila,P. acanthamoebae, E. lausannensis, and P. aeruginosa ATCC 27853 havebeen extracted with a Promega Wizard SV genomic DNA kit (PromegaCorporation, Madison, WI) by following the manufacturer’s instructions.The following primers were used: for W. chondrophila catalase, Wch_KatA_F (5=-CAC CAA AAG AGA TCG CCC AGC CACT-3=) and Wch_KatA_R (5=-TTA GGA GGA ACA GCC TGC TGC TTT TTT GAT TCGC-3=; P. acanthamoebae, Pac_katA_F (5=-CAC CGA GAA TAA AGA TACGCT GAC CAC CA-3=) and Pac_KatA_R (5=-TCA GTT TTT ACG AGAGAG TAG GGCA-3=); E. lausannensis, Ela_KatA_F (5=-CAC CAC AGATAA GCC CCC CCT AT-3=) and Ela_KatA_R (5=-CTA TTT TTT TCTCTT ATC CAG CGC TT-3=); P. aeruginosa, Pae_KatA_F (5=-CAC CGAAGA GAA GAC CCG CCT-3=) and Pae_KatA_R (5=-TCA GTC CAG CTTCAG GCC GAG-3=). The following PCR conditions were used for geneamplifications: initial incubation at 98°C for 30 s, followed by 35 cycles of98°C for 10 s, 68°C for 30 s, and 72°C for 90 s, and an extension of 10 minat 72°C. The annealing temperature was lowered to 61°C for E. lausannen-sis and P. acanthamoebae catalase and to 60°C for P. aeruginosa. High-fidelity Phusion (Fermentas)-amplified genes were cloned in pET200TOPO vector (Invitrogen). The expression of protein was done inBL21Star (Invitrogen). Purification of protein was performed in nonde-naturing conditions with MagneHis (Promega). Purified proteins wereconcentrated in phosphate-buffered saline (PBS) with Amicon columnsaccording to the manufacturer’s instructions (Millipore). Protein concen-trations were determined by fluorescence with a Qubit protein assay byfollowing the manufacturer’s instructions (Invitrogen).

Enzymatic activity. Catalase activity was assessed by decrease in ab-sorption of H2O2 at 240 nm due to degradation as described by Li andSchellhorn, with minor modifications (41). Briefly, hydrogen peroxidewas used at a concentration of 10 mM. Purified proteins were used at aconcentration of 50, 200, or 500 ng depending on the enzymatic activity.PureGrade 96-well plates (Brand, England) were used in a SynergyH1microplate reader (BioTek) at 240 nm.

Elementary bodies of P. acanthamoebae, E. lausannensis, W. chondro-phila, and S. negevensis grown in amoebae were purified according toBertelli et al. (16), with a minor modification. Prior to gastrographingradient ultracentrifugation, the bacteria were treated for 1 h at 37°C withDNase to reduce amoebal protein contamination. An aliquot of frozenbacteria then was washed twice with PBS before exposure to hydrogenperoxide. The bacterial concentration was determined by quantitativePCR (qPCR) with the pan-Chlamydiales primer pair and probe (42). A P.aeruginosa (ATCC 27853) overnight culture was washed twice with PBSand pelleted for 10 min at 8,000 � g at room temperature before exposureto hydrogen peroxide. Bacterial concentration was assessed by determin-ing CFU counts. A. castellanii liquid axenic cultures were counted in akovar chamber, and 106 amoebae were pelleted at 500 � g for 5 min andwashed twice with PBS prior to exposure to hydrogen peroxide.

Nucleotide sequence accession numbers. Newly determined chla-mydial catalase sequences were deposited in GenBank under accessionnumbers KC514601 to KC514620 (see Table S1 in the supplemental ma-terial).

RESULTSGenetic and phylogenetic analyses of chlamydial catalases. Aprototypic small-subunit catalase from Staphylococcus aureus wasused to perform a BLASTP search on all Chlamydiales genomesavailable, including unpublished, in-house, unfinished genomesof Criblamydia sequanensis, Estrella lausannensis, Protochlamydianaegleriophila, and Neochlamydia hartmannellae. Catalases werefound in two members of the Criblamydiaceae family (C. se-quanensis [KatE] and E. lausannensis [KatA]) and in two membersof the Parachlamydiaceae family (P. acanthamoebae [KatA] and N.hartmannellae [KatE]), as well as the previously annotated KatA ofWaddlia chondrophila. When using the catalase sequences of all ofthese members of the Chlamydiales order to perform additionalBLASTP and PSIBLAST searches, no additional catalases wereidentified in any of the available genomes of Chlamydiaceae, Sim-kaniaceae, or Protochlamydia spp.

The amino acid sequence identity of the Chlamydia-relatedbacterial proteins compared to Staphylococcus aureus KatA and tothe prototypical small-subunit Pseudomonas aeruginosa KatAranged between 35 to 60% and 38 to 64%, respectively. Comparedto catalases from clade 2, the identity to KatE of C. sequanensisreached 45 to 68%, except to KatE of N. hartmannellae (75.9%)(see Fig. S1A in the supplemental material). Identity among mem-bers of clade 3 reached around 54% sequence similarity, that be-tween KatA of E. lausannensis and KatA of P. acanthamoebae was77.6%, and that between KatA of W. chondrophila and L. long-beacheae was 77% (see Fig. S1B). These differences in identity ledus to perform a detailed phylogenetic analysis of the chlamydialcatalase proteins.

In addition to reference sequences used by Klotz et al. andZamocky et al., we added the monofunctional catalases found inother amoeba-resisting bacteria (Legionellales, Mycobacteria, andBradyrhizobiales) (12, 43, 44). Moreover, catalases of amoebal or-igin were added to determine a possible lateral gene transfer fromthe host. Sequences were directly retrieved by BLASTP fromamoebal genomes available from NCBI (Dictyosteliida, Acan-thamoeba, Tetrahymena, and Naegleria). For the amoebae belong-ing to the Jakobidae family, the sequences were reconstructedfrom expressed sequence tags (ESTs) retrieved by tBLASTn, sinceno genome was available at the time. Other amoebal sequencesfound in the EST database did not cover the whole protein se-quence; therefore, they were excluded. Moreover, the three high-est BLAST hits for each chlamydial catalase was added to the align-ment as well. Phylogeny of the bacterial catalases was performedwith maximum likelihood (ML), maximum parsimony (MP), andMrBayes (MB), all resulting in clear separation of the three cata-lase categories (i.e., clade 1, 2, and 3) (Fig. 1; also see Fig. S2 in thesupplemental material). Sampled trees from MB analyses showedconstant posterior probabilities, excluding any problem with con-vergence (see Fig. S1C).

Tree topology was conserved between ML and MB in clade 1and clade 2, although with lower confidence in some nodes of ML(see Fig. S2A in the supplemental material). For MP, the Bacillalesbranch of clade 2 catalases clustered with Mycobacteria instead ofbranching off after E. coli, as occurs in the two other analyses.

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However, the phylogenetic relationships of Bacillales could not beaccurately determined, since the bootstrap support was lowerthan 0.5 (see Fig. S2B). The catalases present in C. sequanensis andN. hartmannellae were assigned to the clade 2 catalases that com-prise large tetrameric catalases. As previously observed, the neg-ibacteria catalases are divided into 2 branches (12). KatE of C.sequanensis and KatE of N. hartmannellae clustered with thebranch comprising the posibacterial lineage (Fig. 1A).

Amoebal clade 3 catalases all clustered together with a deeprooting compared to other bacterial clade 3 catalases. There weresmall differences in node placement between the three methods.This was mostly due to the inability of MP and ML to infer thephylogenetic relationship of a few sequences, as can be seen by thelow bootstrap values (see Fig. S2A and B in the supplemental ma-terial). The chlamydial clade 3 catalases separate into two distinctbranches with all three methods. Within the W. chondrophila KatAbranch, five nodes had very low values for both MP and ML (seeFig. S2A and B). However, with MB, the phylogenetic relationshipwas determined with high posterior probabilities (Fig. 1A). E. lau-sannensis and P. acanthamoebae cluster in the same branch, butnot together.

To better understand the origins of these different catalases inChlamydiales, we analyzed their genetic environment and ob-served seven transposase and integrase elements immediately up-stream of katA of W. chondrophila (Fig. 1B), as well as seven trans-

posase and integrase elements 15 genes downstream of waddlialkatA. None of the genes located between these transposases exhib-ited a BLASTP hit with an L. longbeacheae gene, precluding con-firmation that katA was acquired from L. longbeacheae. However,the presence of these mobile elements supports the hypothesis of ahorizontal acquisition of katA by W. chondrophila. No trans-posases were identified around the other catalases. Moreover, thegenetic environment of the KatA-encoding gene from E. lausan-nensis and P. acanthamoebae was not conserved, despite these pro-teins exhibiting a sequence similarity of 78% and clustering to-gether in phylogenetic trees. However, the synteny is low betweenmembers of different families of the Chlamydiales order (45), ex-plaining the absence of the conserved genetic environment despitea likely common origin for E. lausannensis and P. acanthamoebaekatA.

Conservation of domains and motifs. The heme binding sitesare conserved among all clades of monofunctional catalases (43).Therefore, we analyzed the sequences of the chlamydial catalasesto determine motif conservation. Differences in prevalence of agiven amino acid within the motifs were observed depending onthe catalase clade. Amino acid variants that were mostly found inclade 2 catalases are highlighted in boldface in Fig. 2. The proximalheme-binding site was very conserved and was detected in all ofthe chlamydial catalases (Fig. 2E). The active sites in both siteswere conserved (Fig. 2D and E, red boxes). For the distal heme

FIG 1 Gene environment and phylogenetic tree of typical bacterial catalases, including chlamydial catalases. (A) Representative tree of bacterial catalases,obtained from http://peroxibase.toulouse.inra.fr/, with catalase sequences of Chlamydiales and other amoeba-resisting bacteria. The evolutionary history wasinferred by using MB (1,000,000 generations) and confirmed by ML (500 bootstraps) and MP (500 bootstraps). Only posterior probabilities below 1 are marked(red, 0.99 to 0.95; yellow, 0.95 to 0.9; green, 0.85 to 0.8; blue, 0.77 to 0.76). The bar indicates 0.1 amino acid substitutions per site. (B) The gene environment ofthe catalases present in the different Chlamydiales members exhibited no homology. Only W. chondrophila encodes transposases and integrases near the catalaselocation, suggesting that this catalase was obtained by lateral gene transfer.

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binding site (Fig. 2D), analysis by PROSITE did not give any hitfor KatA of P. acanthamoebae and KatA of E. lausannensis. Thiswas due to two mutations at positions F44Q and S59W, respec-tively (Fig. 2D). The phenylalanine is very conserved, and its re-placement with glutamine could strongly affect the binding site,especially its topology. The second substitution is also quite prob-lematic, since tryptophan is more reactive and bulkier than serine.

Moreover, the sequences were analyzed for the conservation ofsurface epitopes determined in a previous study (46). Since theconsensus sequence for these epitopes was determined with onlyfour sequences from small-subunit catalases, we further con-firmed them by analyzing the MUSCLE alignment of all bacterialcatalases obtained from http://peroxibase.toulouse.inra.fr/ (seeFig. S3 in the supplemental material). All three epitopes were con-served in all clades of catalases, with minor differences betweensmall- and large-subunit catalases, except for the second epitope.The second epitope had a more conserved sequence in clade 2catalases than clade 3. Moreover, the consensus sequences differedsubstantially between the two clades (see Fig. S3, orange, green,and blue boxes). The first epitope (Fig. 2A) was conserved in allChlamydiales catalases. The last surface epitope (Fig. 2C) was wellconserved in all Chlamydiae members except KatA of W. chondro-phila, which presented two mutations in the more conserved sites(N304D and F306H). Moreover, at position 305, KatA of W. chon-drophila had a phenylalanine, like large-subunit catalases.

Enzymatic activity. Since Chlamydiales present two develop-mental stages, the activity of the catalase would be required duringthe early steps of infection, when the bacteria are still in theirelementary body form. The amount of bacteria present in each vialwas determined by qPCR with the pan-Chlamydiales primers. Atotal of 107 purified elementary bodies of P. acanthamoebae, E.lausannensis, C. sequanensis, N. hartmannellae, and W. chondro-phila, all encoding a catalase, were exposed to 0.2 M hydrogenperoxide (Fig. 3A). All of them produced oxygen bubbles to anextent similar to that of the P. aeruginosa positive control. How-ever, N. hartmannellae and W. chondrophila displayed reducedoxygen bubble formation. A mock negative control of the amoebalcoculture was also tested and proved to be negative. Simkanianegevensis was used as a second negative control, since it does notencode a catalase. Therefore, as expected, it only produced one

small oxygen bubble. This could be due to a slight contaminationof purified elementary bodies by Acanthamoebae castellanii cata-lase. However, the contamination is insignificant compared to thecatalase activity displayed by the different bacteria. The clade 3catalase of A. castellanii was also blocked by 0.1 M azide. Theactivity of the catalase was blocked in all bacteria with 0.1 M azide,a noncompetitive inhibitor of catalases (47). Only when the con-centration was reduced to 1 mM did we again observe oxygenformation for P. aeruginosa bacteria, while E. lausannensis showedno catalase activity even at 0.1 mM azide (Fig. 3A).

Once we observed this oxygen production, we determined thecatalytic activity of these catalases at various pHs in order to definewhether the activity correlates with host range and phagolysosomesurvival. As a positive control, we cloned KatA from P. aeruginosa.To control for eventual contamination by the Escherichia coli cat-alase, we used as a negative control Hsp60 from W. chondrophilapurified under the same conditions as the catalases. The enzymaticactivity was assessed by measuring the absorption of H2O2. ForKatA of P. acanthamoebae and KatA of W. chondrophila, we had toincrease the amount of protein from 200 ng/ml to 2 �g/ml and 800ng/ml, respectively, to detect a sustained degradation of hydrogenperoxide (Fig. 3B). The catalytic units (Fig. 3C) derived from theassay differed quite substantially depending on the species. KatAof P. acanthamoebae had the least ability to degrade hydrogenperoxide. KatA of E. lausannensis was more efficient at pH 7.0,whereas KatA of W. chondrophila appeared to be less sensitive tochanges in pH.

In silico modeling of chlamydial catalases. Several crystalstructures exist for bacterial clade 2 and 3 catalases. Therefore, weused this information to build an in silico model of the chlamydialcatalases using 3DJIGSAW (28–30) and SWISS-MODEL Work-space (31–34). The tetrameric structure of the clade 3 catalases wasbuilt on the crystal structure of Enterobacter faecalis (1SI8) (seeFig. S4A in the supplemental material) for all three chlamydialcatalases. The organization of the tetramer was retained for allchlamydial small-subunit catalases (see Fig. S4A). For the chla-mydial large-subunit catalases, the crystal structure of E. coli HPII(1IPH) was used (see Fig. S4B). The organization of the tetramerof KatE of N. hartmannellae catalase was more similar to HPIIfrom E. coli than KatE of C. sequanensis (see Fig. S4B).

FIG 2 Surface epitopes and heme binding domains. Amino acids in boldface are differently conserved in large-subunit catalases. (A to C) Surface epitopesconserved in Chlamydiales, previously determined by in silico modeling of H. hepaticus (46); note the N304D and F306H mutations (arrows; Pseudomonasnumbering). (D) Distal site of the prosthetic heme with catalytic histidine (red box); note the F44Q and S59W mutations (arrows). (E) Proximal site of theprosthetic heme with catalytic tyrosine (red box).

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To determine the conservation of the tertiary structure of thetetramer subunits, the RMS values for each chlamydial catalasewere determined. Overall, the tertiary structures were well con-served, with only limited deviations in the loop regions (Fig. 4A toC; also see Fig. S4C in the supplemental material). KatE of C.sequanensis presented one helix with more than a 5-Å deviationfrom the crystal structure. However, this part of the subunit isinvolved in neither the contact site of the tetramer nor in thecatalytic domain (see Fig. S4C, left). Moreover, the quality of thetertiary structure was further determined by looking at residueswith bad backbone conformation and side chains without hydro-gen bonds. There were 24 residues in the C terminus with clashesin the backbone. Of these 24, 15 are mutated compared to E. coliHPII and 4 were not aligned. The remaining residues were close tomutated residues that caused the distortion of the backbone (seeFig. S5A). Moreover, in the C terminal of KatE of C. sequanensis,nine residues were mutated to prolines that strongly influenced

the backbone orientation. Only three prolines (P383, P522, andP543) were not in an optimal conformation. As for KatE of C.sequanensis, most of the residues (11) that clashed with the back-bone were located in the C-terminal part of the protein.

Chlamydial small-subunit catalases had no side chains thatlacked hydrogen bonds. In KatA of P. acanthamoebae, KatA of E.lausannensis, and KatA of W. chondrophila there were only fiveresidues, of which 3 were mutated compared to the Helicobacterpylori (2IQF) catalase sequence (see Fig. S4B in the supplementalmaterial). The mutations were not all located at the same residues.Moreover, KatA of P. acanthamoebae catalase had one prolinemutation (Q379P), KatA of W. chondrophila had two (V6P andV55P), and KatA of E. lausannensis had one (V6P) that affectedthe backbone organization.

The water molecules for both catalases were added to the cat-alytic site residues according to Diaz et al. (48) (Fig. 4A). For thesmall-subunit catalase, the residue orientation is almost com-

FIG 3 Enzymatic activity of chlamydial catalases in vitro. (A) Elementary bodies (EBs; 107) of Chlamydiales and P. aeruginosa were exposed to 0.2 M H2O2.Catalase activity is blocked by 0.1 M azide. (B) Degradation of hydrogen peroxide by purified catalase was followed by a decrease in absorbance at 240 nm.Catalase of P. acanthamoebae exhibited the weakest activity. The activity of the enzymes was conserved even when the pH was lower. Results are means from atleast six independent experiments with standard errors of the means for each condition and protein. Hsp60 of W. chondrophila was used as a negative control. (C)Catalytic units derived from a decrease in absorbance are in M�1 · s�1.

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pletely conserved. For KatA of W. chondrophila R53, the side chainis in another orientation that replaces the hydrogen bond with theheme with the water molecule (W2) (Fig. 4B). The rotation ofN128 in KatA of P. acanthamoebae and KatA of E. lausannensiscaused a weakening of the interaction with the water molecule (3.2Å) of the nitrogen of the histidine ring and the oxygen of theasparagine side chain (Fig. 4C). Moreover, the interaction mostlikely occurs with the oxygen and not the nitrogen of the aspara-gine side chain, like in H. pylori. In KatA of P. acanthamoebae, theslight tilt of H53 further increased the distance to the water mol-ecule, weakening the interaction. All other hydrogen bonds werein the range of moderate interactions (2.5 to 3.2 Å). The flipping ofthe H198 side chain in KatA of P. acanthamoebae disrupted thehydrogen bond chain between the catalytic residues Y338, R334,H198, and N328. Indeed, the distances passing 4 Å did not allowfor hydrogen bond formation. This could affect the stability of thecatalytic site. For chlamydial clade 2 catalases, the catalytic site wasalso conserved, and all hydrogen bonds could be formed (see Fig.S4D in the supplemental material). Only Y415 in KatE of C. se-quanensis had a 14° shift of the aromatic ring, lengthening thedistance between the tyrosine and the iron of the heme from 2.11to 2.35 Å. However, the tyrosine was still close enough to interactwith the iron.

DISCUSSION

In this work, we identified catalase-encoding genes in the genomesof five different Chlamydia-related bacteria, and we could demon-strate their enzymatic activity and a significant conservation oftheir active sites. These catalases belonged to clade 2 (KatE of N.hartmannellae and KatE of C. sequanensis) and clade 3 (KatA of W.chondrophila, KatA of E. lausannensis, and KatA of P. acantham-oebae) catalases. Moreover, waddlial catalase was branching farfrom the other two clade 3 catalases, encoded by P. acanthamoebaeand E. lausannensis, suggesting a complex evolutionary history ofthese proteins. The limited amount of sequences available forchlamydial catalases did not allow us to determine definitelywhich one is most likely to be the ancestral Chlamydiales catalase(Fig. 5A). However, according to phylogenetic analyses per-formed by Klotz et al., the first catalase was likely a large-subunitcatalase that underwent gene duplication with loss of the C-termi-nal domain and partial loss at the N-terminal domain (12). Thus,it is likely that the common ancestor of the Chlamydiales order hada large-subunit catalase that was lost by most of the species. More-over, lateral gene transfer of large-subunit catalases was only ob-served from bacteria to archaea and not among bacteria (12).

The clade 3 catalase shared by E. lausannensis and P. acantham-

FIG 4 In silico modeling of small-subunit chlamydial catalases. Tertiary structure of small-subunit chlamydial catalases marked according to RMS values. (A) W.chondrophila KatA tertiary structure is strongly conserved. (B and C) Tertiary structures of KatA of P. acanthamoebae and KatA of E. lausannensis are conserved,except in some loops (arrows). (D) Catalytic site of H. pylori (white; PDB code 2IQF), P. mirabilis (salmon; PDB code 1M85), and E. faecalis (yellow, PDB code1SI8). Numbering is according to that for H. pylori. (E) Catalytic site of W. chondrophila KatA (purple). All residues are conserved, except R53 (arrow). (F)Catalytic site of KatA of P. acanthamoebae (red) and KatA of E. lausannensis (white). The orientation of the active-site residues is conserved, except for H198 ofP. acanthamoebae (arrow) and N128 in both bacteria (arrow). Numbering is according to that of KatA of the P. acanthamoebae sequence. Water molecules aremarked W1 to W3, with hydrogen bonds shown in dotted green lines. The heme of H. pylori was introduced into the model structure of the small-subunitchlamydial catalases.

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oebae was probably acquired later, since this enzymatic clade orig-inated later in posibacteria and was then distributed to other spe-cies by lateral gene transfer (12). This transfer in Chlamydialeslikely has taken place after the separation of the Criblamydiaceae/Parachlamydiaceae ancestor from the Waddliaceae, since they en-code clade 3 catalases of different origins (Fig. 5A). Since E. laus-annensis and P. acanthamoebae were clustering in the same branchbut not together, it is more likely that they both received the cat-alase from a Rhizobiales member on a different occasion than aninternal transfer from one to the other after lateral gene transferfrom a nonchlamydial bacteria (Fig. 5B). The transfer of the cat-alase present in E. lausannensis and P. acanthamoebae probablyoccurred from a Rhizobiales precursor, since the majority of thefirst 30 BLASTP hits belong to this order. For W. chondrophila, theorigin of the lateral gene transfer is probably an Actinomycetales orLegionellales isolate rather than one of the Planctomycetes BLASTbest hits. This hypothesis is based on the fact that Legionellalesisolates and several members of the Actinomycetales are able tosurvive within amoebae (44). Interestingly, host-associated gen-era, like Brucella and Bordetella, lost the large-subunit catalaseupon acquisition of the clade 3 catalase. Such a loss could explainthe absence of the clade 2 catalase from the chlamydia-relatedbacteria that encode a clade 3 catalase. As proposed by Klotz et al.,this might be due to the presence of clade 3 catalases in eukaryotesthat are forcing the bacteria to adapt to the physiological selectivepressure present within the host (12). When looking at the mostcommon human bacterial pathogens, the majority encode a cata-lase or a catalase-peroxidase. As seen for other anaerobic bacteria,Streptococcaceae do not possess a catalase, since their exposure toreactive oxygen species is much lower. Mycoplasma, Rickettsiarickettsii, Borrelia burgdorferi, Treponema pallidum, and Chlamy-diaceae all underwent strong genome size reduction and adapta-tion to a specific niche, making the presence of a catalase redun-dant. Since the reservoir of Chlamydia-related bacteria is muchbroader, these bacteria need a more diverse panel of virulencefactors to adapt to each host. For W. chondrophila and E. lausan-

nensis, the presence of a catalase could indeed prove useful duringthe infection of humans when encountering professional phago-cytes. Even though the members of the Chlamydiales order allshare the same replicative cycle, the composition of the bacterialinclusion varies considerably between the Chlamydiaceae andChlamydia-related bacteria. The inclusion of Chlamydiaceae es-capes the endocytic pathway early and associates with Golgi pro-teins and membrane (49–51). In macrophages, W. chondrophila isonly transiently associated with the endocytic pathway but avoidslysosomal fusion, as it is associated with the endoplasmic reticu-lum (8). Conversely, in macrophages, P. acanthamoebae does ac-quire LAMP-1 but prevents further maturation of the bacterialinclusion into a digestive lysosome (52). Perhaps due to thesedifferences in the early intracellular trafficking of the Chlamydia-les, W. chondrophila and P. acanthamoebae require a catalase tocounteract initial exposure to ROS, whereas Chlamydiaceae donot. A better understanding of the trafficking of other Chlamydia-related bacteria, like Protochlamydia naegleriophila, which doesnot encode a catalase, and C. sequanensis, which encodes a clade 2catalase, is essential to define the roles of these enzymes.

Despite significant differences at the amino acid sequence level,tertiary structure modeling showed significant conservation, es-pecially around the catalytic sites. Histidine 198 of KatA of P.acanthamoebae probably destabilizes the interaction between theheme and the catalytic tyrosine due to a disruption of the hydro-gen bond chain. This correlates with the reduced enzymatic activ-ity of KatA of P. acanthamoebae observed in vitro. Such loweractivity of KatA of P. acanthamoebae compared to that of KatA ofW. chondrophila may affect the ability of these bacteria to growwithin macrophages, since a rapid response to ROS production iscrucial to prevent damage and activation of inflammatory signal-ing pathways. Indeed, W. chondrophila has a productive and rapidgrowth cycle in macrophages (7, 8), whereas P. acanthamoebaegrows only poorly in macrophages (10, 11); this might partially bedue to their different fates in early steps of intracellular trafficking.However, the presence of a catalase is not essential for growth in

FIG 5 Model of catalase acquisition and loss in Chlamydiales. (A) MrBayes phylogeny of fully sequenced Chlamydiales based on concatenation of GyrA, GyrB,RpoA, RpoB, RecA, SecY, TufA, and TopA amino acid sequences. Only nodes with posterior probability inferior to 1 are marked. The tree constructed with MLplaced the Criblamydiaceae node before that for Waddliaceae. Families are delimited by colored boxes. Distribution of catalases in the Chlamydiales order isdepicted in the column. (B) Two hypotheses for lateral transfer of clade 3 catalase in P. acanthamoebae and E. lausannensis. Transfer from a Rhizobiales memberto one of the two chlamydial species and then internal transfer (1) or transfer from a Rhizobiales member to both species at about the same time (2). (C) Catalaseof W. chondrophila could originate from a lateral transfer from Legionellales (1) or an Actinomycetales isolate (2).

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free-living amoebae, since Protochlamydia spp. do not encode anycatalases but nevertheless grow successfully in these protists (53,54). As revealed by phylogenetic analyses, the horizontal transferof clade 3 catalases probably occurred on several occasions andafter the divergence of the different families within the Chlamyd-ia-related bacterial branch. Moreover, within the Parachlamydi-aceae, the Protochlamydia spp. that do not encode any catalasediverged prior to acquisition of catalases by Parachlamydia spp.

ACKNOWLEDGMENTS

This work was supported by the Swiss National Science Foundation (proj-ect no. PDFMP3-127302). B.R. is supported by the Swiss National ScienceFoundation within the PRODOC program “Infection and Immunity.”

We declare no conflicts of interest.We thank C. Bertelli and T. Pillonel for helpful comments.

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