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RESEARCH ARTICLE

crossm Role of Acetyltransferase PG1842 in Gingipain Biogenesis in Porphyromonas gingivalis Arunima Mishra,a Francis Roy,a Yuetan Dou,a Kangling Zhang,b Hui Tang,b Hansel M. Fletchera,c

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a

Division of Microbiology and Molecular Genetics, Department of Basic Sciences, School of Medicine, Loma Linda University, Loma Linda, California, USA

b

Department of Pharmacology, University of Texas Medical Branch, Galveston, Texas, USA

c

Institute of Oral Biology, Kyung Hee University, Seoul, Republic of Korea

ABSTRACT Porphyromonas gingivalis, the major etiologic agent in adult periodontitis, produces large amounts of proteases that are important for its survival and pathogenesis. The activation/maturation of gingipains, the major proteases, in P. gingivalis involves a complex network of processes which are not yet fully understood. VimA, a putative acetyltransferase and virulence-modulating protein in P. gingivalis, is known to be involved in gingipain biogenesis. P. gingivalis FLL92, a vimA-defective isogenic mutant (vimA::ermF-ermAM) showed late-onset gingipain activity at stationary phase, indicating the likelihood of a complementary functional VimA homolog in that growth phase. This study aimed to identify a functional homolog(s) that may activate the gingipains in the absence of VimA at stationary phase. A bioinformatics analysis showed five putative GCN5-related N-acetyltransferases (GNAT) encoded in the P. gingivalis genome that are structurally related to VimA. Allelic exchange mutagenesis was used to make deletion mutants for these acetyltransferases in the P. gingivalis vimAdefective mutant FLL102 (ΔvimA::ermF) genetic background. One of the mutants, designated P. gingivalis FLL126 (ΔvimA-ΔPG1842), did not show any late-onset gingipain activity at stationary phase compared to that of the parent strain P. gingivalis FLL102. A Western blot analysis of stationary-phase extracellular fractions with antigingipain antibodies showed immunoreactive bands that were similar in size to those for the progingipain species present only in the ΔvimA-ΔPG1842 isogenic mutant. Both recombinant VimA and PG1842 proteins acetylated Y230, K247, and K248 residues in the pro-RgpB substrate. Collectively, these findings indicate that PG1842 may play a significant role in the activation/maturation of gingipains in P. gingivalis. IMPORTANCE Gingipain proteases are key virulence factors secreted by Porphyromo-

nas gingivalis that cause periodontal tissue damage and the degradation of the host immune system proteins. Gingipains are translated as an inactive zymogen to restrict intracellular proteolytic activity before secretion. Posttranslational processing converts the inactive proenzyme to a catalytically active protease. Gingipain biogenesis, including its secretion and activation, is a complex process which is still not fully understood. One recent study identified acetylated lysine residues in the three gingipains RgpA, RgpB, and Kgp, thus indicating a role for acetylation in gingipain biogenesis. Here, we show that the acetyltransferases VimA and PG1842 can acetylate the proRgpB gingipain species. These findings further indicate that acetylation is a potential mechanism in the gingipain activation/maturation pathway in P. gingivalis. KEYWORDS acetyltransferase, gingipain, Porphyromonas gingivalis

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orphyromonas gingivalis is a well-established, Gram-negative anaerobic oral bacterium involved in chronic periodontitis (1). After dental caries, periodontal diseases are the second most frequent oral diseases, affecting up to 90% of the global popuDecember 2018 Volume 200 Issue 24 e00385-18

Journal of Bacteriology

Received 27 June 2018 Accepted 20 September 2018 Accepted manuscript posted online 24 September 2018 Citation Mishra A, Roy F, Dou Y, Zhang K, Tang H, Fletcher HM. 2018. Role of acetyltransferase PG1842 in gingipain biogenesis in Porphyromonas gingivalis. J Bacteriol 200:e00385-18. https://doi.org/10.1128/JB .00385-18. Editor Victor J. DiRita, Michigan State University Copyright © 2018 American Society for Microbiology. All Rights Reserved. Address correspondence to Hansel M. Fletcher, hfl[email protected].

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lation and posing a major threat to public health (2). Periodontitis is a complex inflammatory disease characterized by bacterial colonization of the gingival sulcus and periodontal pocket, which can result in deepening of the pocket, alveolar bone loss, and in severe cases, tooth loss (3). P. gingivalis is also associated with systemic diseases, such as the initiation and/or progression of cardiovascular disease and rheumatoid arthritis (4, 5). Although several virulence factors are known in the pathogenicity of P. gingivalis, including fimbriae, hemagglutinin, capsule, and lipopolysaccharide, the major virulence factor is a family of cysteine proteases, called gingipains, consisting of an arginine-specific protease (Rgp) and lysine-specific protease (Kgp) (6). The Rgp is encoded by two genes, rgpA and rgpB, while Kgp is encoded by a single gene, kgp (7). These proteases are both extracellular and cell associated. Indeed, approximately 85% of the total extracellular protease activity of P. gingivalis is from gingipains secreted into the extracellular host environment (8). Gingipains are involved in variety of functions needed for the survival of the bacterium in the anaerobic host environment, including the acquisition of essential nutrients, the invasion of host tissues, the inactivation of cytokines and their receptors, and the attenuation of neutrophil antibacterial activities (8, 9). The activity of gingipains must be regulated inside P. gingivalis to inhibit unwanted intracellular proteolytic activity before being secreted into the extracellular environment (10). Therefore, gingipains are translated as inactive proenzymes which then undergo posttranslational processing to generate mature active enzymes (11). For example, RgpB is synthesized as a proenzyme (pro-RgpB) possessing an N-terminal signal peptide, a prodomain, a catalytic domain, and a C-terminal domain (CTD) (12). The maturation of the inactive pro-RgpB to the catalytically active RgpB is complex, with multiple processing steps which are not yet fully defined. Once the pro-RgpB translocates across the inner membrane via the Sec machinery, the N-terminal prodomain is sequentially processed to activate the proenzyme (11). At the same time, the CTD targets the maturing protein to a type IX secretory system, which translocates the maturing RgpB through the outer membrane (13). During the outer membrane translocation process, the CTD of RgpB is removed by a cysteine protease, PG0026 (14), to produce either the mature 48-kDa soluble form or the 70- to 90-kDa highly glycosylated membrane attached form (15). Protein acetylation has emerged as a universal posttranslational modification mechanism in both eukaryotes and prokaryotes (16–19). This protein modification is finely tuned via both enzymatic (by protein acetyltransferases) and nonenzymatic (by metabolic intermediates such as acetyl phosphate) mechanisms (17, 20, 21). In bacteria, acetylation is mainly catalyzed by a specific acetyltransferase enzyme using acetyl coenzyme A (acetyl-CoA) as a donor (22). Protein acetylation has been shown to play a role in bacterial chemotaxis, central metabolism, DNA replication, and bacterial virulence (19, 23–26). Lysine acetylation plays an important regulatory role by changing the biochemical characteristics of proteins, such as their charge, stability, and interactions with other molecules (19, 26). In a recent P. gingivalis acetylome study, Butler et al. identified 92 lysine-acetylated proteins, including the three gingipains RgpA, RgpB, and Kgp (27). This suggests that acetylation is an important posttranslational modification required for gingipain activation. The P. gingivalis vimA gene is part of the bcp-recA-vimA-vimE-vimF-PG0885-PG0886 operon and encodes a putative acetyltransferase protein (28). Previously, we reported reduced gingipain activity in a vimAdefective mutant, FLL92 (29, 30). Compared to that in the W83 wild type, 90% reductions were observed for both Rgp and Kgp activities in FLL92 during the exponential growth phase. However, these activities were increased to roughly 60% of that of the wild-type strain during stationary phase (30). This unique late-onset activity suggests that during exponential phase, gingipain activity in P. gingivalis is regulated in a vimA-dependent manner. However, at stationary phase, the activity may be due to a functional homolog of VimA that is upregulated or activated to complement VimA’s function. In this study, we identified PG1842 as the functional homolog of VimA at stationary phase and showed that both VimA and PG1842 can use pro-RgpB as a jb.asm.org 2

Role of Acetyltransferase PG1842 in P. gingivalis



FIG 1 Quantitative real-time PCR showing fold changed of genes vimE, vimF, PG0885, and PG0886 in vimA mutants FLL92 and FLL102 compared to that in the W83 wild-type strain. In FLL92, all four genes were expressed at low levels, with the least expressed gene, vimF, downregulated more than 6-fold. By contrast, the expression levels of vimE, vimF, PG0885, and PG0886 were unaffected in FLL102. Total RNA from W83, FLL92, and FLL102 strains were extracted from exponentially growing bacterial cells (OD600 of ⬃0.8). Fold change in expression levels was calculated by the 2⫺ΔΔCT method. Gene expression levels were normalized to those of 16S rRNA transcripts. Data represent the averages from three independent experiments. Each experiment was done in triplicates. Error bars represent the standard deviations from the means. **, P ⬍ 0.05; ***, P ⬍ 0.005 versus W83.

substrate for acetylation. The results contribute to an understanding of the gingipain activation/maturation pathway(s) in P. gingivalis. RESULTS Inactivation of the vimA gene has a polar effect on the downstream genes in the operon. P. gingivalis FLL92 (vimA::ermF-ermAM) carries a vimA gene that is inactive due to the insertion of the ermF-ermAM antibiotic resistance cassette containing a transcriptional terminator (29). To clarify the polar effects on the other genes in the transcriptional unit, the expression levels of the downstream genes in P. gingivalis FLL92 were compared to those in the W83 wild-type strain via reverse transcriptionquantitative PCR (qRT-PCR). As shown in Fig. 1, the vimE, vimF, PG0885, and PG0886 genes were downregulated 3.6-, 6.5-, 5.6-, and 2.8-fold, respectively. To further determine the specific effect(s) of the vimA gene on the phenotypic properties of P. gingivalis, an isogenic mutant was created by replacing this gene with an ermF antibiotic resistance cassette that is missing a transcriptional terminator. After the electroporation of P. gingivalis with the purified PCR fusion fragment (see Materials and Methods), several nonpigmented erythromycin-resistant colonies were detected after 7 days of incubation. The replacement of the vimA gene with the ermF cassette in these mutants was confirmed by PCR and DNA sequencing (data not shown). One randomly chosen mutant, designated FLL102 (ΔvimA::ermF) was selected for further studies. To assess any polar effect of the vimA gene deletion in P. gingivalis FLL102, the expression levels of the vimE, vimF, PG0885, and PG0886 genes were determined by qRT-PCR. In contrast to that observed in FLL92, the expression levels of these genes were unaffected in FLL102 (Fig. 1). P. gingivalis FLL102 was used in the remainder of our studies to evaluate gingipain biogenesis. P. gingivalis FLL102 exhibits late-onset gingipain activity. During the exponential growth phase, gingipain activity in P. gingivalis FLL102 (ΔvimA::ermF) was reduced more than 90% compared to that in the W83 wild type (Fig. 2A). At stationary phase, the gingipain activity of the vimA deletion mutant was increased in contrast to the activity in W83, which remained the same at both exponential and stationary growth phases. Compared with that at exponential phase, the Rgp and Kgp activities were increased to 23% and 40%, respectively (Fig. 2B). Altogether, the data suggest that under the same physiological conditions, there is a late onset of proteolytic activity in the P. gingivalis FLL102 isogenic mutant compared to that in W83. VimA is a putative acetyltransferase that belongs to the GNAT family. A bioinformatics analysis of the amino acid sequence of the VimA protein predicted an December 2018 Volume 200 Issue 24 e00385-18

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FIG 2 Gingipain activity of the W83 wild type and the ΔvimA mutant and its ΔPG1088, ΔPG1254, ΔPG1358, ΔPG1761 and ΔPG1842 isogenic derivatives at exponential (A) and stationary (B) growth phases. P. gingivalis strains were grown in BHI medium supplemented with vitamin K and hemin. Activities against Rgp or Kgp were tested in whole-cell cultures. The gingipain activities were normalized to that of W83, set as 100%, and the mutants are reported as a percentage thereof. Experiments were carried out in three independent repeats. Error bars represent the standard deviations from the means. ***, P ⬍ 0.005 versus W83. Compared to those in W83, both Rgp and Kgp activities were reduced more than 90% in the ΔvimA mutant (A), which showed late-onset Rgp (23%) and Kgp (40%) activity at stationary phase (B). The ΔvimA-ΔPG1842 double mutant did not show any late-onset activity at stationary phase.

acetyltransferase domain of the GCN5-related N-acetyltransferase (GNAT) family (Fig. 3A). Orthologs of vimA are present in many anaerobic bacteria, such as Tannerella forsythia, Clostridium botulinum, Rhodobacter sphaeroides, and Parabacteroides distasonis (28). A BLAST analysis against the NCBI protein database using the amino acid sequence of VimA revealed homology to GNAT enzymes from Porphyromonas gulae (92% identity), Porphyromonas crevioricanis (37% identity), T. forsythia (39% identity), Parabacteroides (37% identity), and Geobacillus thermodenitrificans (34% identity). The secondary structure (predicted by I-TASSER) of VimA showed characteristic ␤-strands and ␣-helices (␤1-␣1-␣2-␤2-␤3-␤4-␣3-␤5-␣4-␤6) that are conserved in the GNAT superfamily (Fig. 3B). This structure was similar to the six ␤-strands and four ␣-helices found in the secondary structure (derived from X-ray crystallography) of the GNAT aminoglycoside 3-N-acetyltransferase (AAC-3) from Serratia marcescens (Fig. 3C) (31). It is noteworthy that the primary sequence of VimA did not show any observable sequence identity with the S. marcescens AAC-3. However, both VimA and AAC-3 acetyltransferases displayed the four conserved GNAT C, D, A, and B motifs. Motif A, the core of the GNAT fold, is highly conserved and generally has the R/Q-X-X-G-X-G/A sequence critical for acetyl-CoA recognition and binding (16). Similar to that in AAC-3, the acetyl-CoA binding sequence QIREGQG (conserved residues are underlined) was found in the predicted motif A of VimA (indicated by asterisks in Fig. 3B and C) (31). To ascertain a functional homolog of VimA that can complement the mutation in the ΔvimA mutant at stationary growth phase, the P. gingivalis W83 genome was surveyed for other acetyltransferases. Five genes (PG1088, PG1254, PG1358, PG1761, and PG1842) encoding putative GNAT enzymes were identified. PG1088 (Ac_7; pfam13508), PG1254 (Ac_3; pfam13302), PG1358 (Ac_3; pfam13302), PG1761 (Ac_1; pfam00583), and PG1842 (Ac_9; pfam13527) were predicted to belong to different families in the GNAT superfamily (Fig. 3A) (32). Although the primary sequence of VimA showed very low sequence identity with PG1088, PG1254, PG1358, PG1761, and PG1842, the secondary structures (predicted by I-TASSER) displayed a similar classic GNAT fold with motif A showing the conserved R/Q-X-X-G-X-G/A sequence for acetyl-CoA binding (Fig. 3D). ⌬vimA-⌬PG1842 mutant does not exhibit late-onset gingipain activity at stationary phase. To determine if any of the putative acetyltransferases identified above December 2018 Volume 200 Issue 24 e00385-18

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FIG 3 (A) Predicted domain structures of VimA, PG1088, PG1254, PG1358, PG1761, and PG1842. Conserved domains were predicted by the NCBI CDD database. (B) Secondary structure prediction of VimA (by I-TASSER) showing characteristic ␤-strands and ␣-helices (␤1-␣1-␣2-␤2-␤3-␤4-␣3-␤5-␣4-␤6) of the GNAT superfamily. The conserved sequence QIREGQG in motif A is shown by asterisks. (C) Secondary structural elements of S. marcescens aminoglycoside 3-N-acetyltransferase derived from the X-ray structure. The structure shows the typical GNAT fold, including six ␤-strands and four ␣-helices. The conserved sequence RRQGIG in motif A is shown by asterisks. (D) Secondary structures of PG1088, PG1254, PG1358, PG1761, and PG1842 (predicted by I-TASSER) showing conserved sequence R/Q-X-X-G-X-A/G in motif A (shown by asterisks). The conserved motifs, sequentially labeled C, D, A, and B, are represented with boxes. Residues in ␤-strands are highlighted in gray, whereas residues in ␣-helices are highlighted in cyan.


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could functionally complement the VimA protein, the genes encoding these enzymes were replaced with the tetQ (tetracycline resistance) transcriptional terminatorless cassette in the vimA-defective mutant FLL102 genetic background. The isogenic double mutants generated (ΔvimA-ΔPG1088, ΔvimA-ΔPG1254, ΔvimA-ΔPG1358, ΔvimAΔPG1761, and ΔvimA-ΔPG1842) were confirmed by PCR and DNA sequencing (data not shown). Similar to that in the ΔvimA parent strain, the exponential growth phase cultures of the ΔvimA-ΔPG1088, ΔvimA-ΔPG1254, ΔvimA-ΔPG1358, ΔvimA-ΔPG1761, and ΔvimA-ΔPG1842 double mutants showed little or no proteolytic activity (Fig. 2A). However, at stationary phase, the ΔvimA-ΔPG1842 mutant did not show any late-onset activity in contrast to that of other double mutants, all of which had late gingipain activity similar to that of the ΔvimA isogenic strain (Fig. 2B). These results suggest that PG1842 is a potential functional VimA homolog. No processing of inactive proenzyme gingipain species into active mature forms in the ⌬vimA-⌬PG1842 mutant at stationary phase. The studies described above were extended by comparing the presence of major protease species in the ΔvimA mutant and its ΔvimA-ΔPG1842 isogenic derivative by immunoblot analysis. As shown in Fig. 4A, during exponential phase, RgpB was secreted into the extracellular fraction of the ΔvimA mutant as an ⬃70-kDa proenzyme. This immunoreactive band was not detected at stationary phase; however, a band similar in size to the band representing the catalytic domain in the W83 wild type was observed at stationary phase in the extracellular fraction. In contrast, for the ΔvimA-ΔPG1842 isogenic strain, a 70-kDa proenzyme band was seen in the extracellular fractions during both exponential and stationary growth phases. When the blot was probed with an anti-Kgp antibody (Fig. 4B), the high-molecular-weight inactive forms of Kgp were detected at exponential phase in the ΔvimA mutant. At stationary phase, a band similar in size to the catalytic domain of Kgp in W83 was observed in the extracellular fraction. In the ΔvimA-ΔPG1842 isogenic mutant, both exponential and stationary growth phases were characterized by the presence of only the high-molecular-weight inactive forms of Kgp. An immunoblot analysis with an anti-RgpA antibody showed similar results as those with the anti-Kgp antibody (data not shown). Altogether, these results suggest that in the absence of PG1842 (ΔvimA-ΔPG1842), the inactive proenzyme species were not processed into the mature/active forms at stationary phase. PG1842 is upregulated in the ⌬vimA mutant at stationary phase compared to that in exponential growth phase. Because PG1842 may compensate for the VimA defect at stationary phase, we evaluated its modulation during the exponential and stationary growth phases via qRT-PCR. In the W83 wild-type strain, all five acetyltransferase genes were downregulated at stationary phase. The most downregulated gene was PG1254 (5.5-fold), while the other genes were downregulated between 1.1- and 2.3-fold (Fig. 5A). In contrast to the expression of the PG1254 gene, which was downregulated 1.7-fold, the other acetyltransferase genes were upregulated (between 2.2- and 3-fold) at stationary phase in the ΔvimA mutant (Fig. 5B). Furthermore, the expression levels of PG1088, PG1254, PG1358, PG1761, and PG1842 in the ΔvimA mutant were compared with those in the W83 strain. At exponential phase, all genes were upregulated at low levels, between 1.4- and 1.9-fold (Fig. 5C). In contrast, the genes were highly upregulated at stationary phase (⬃7- to 9-fold upregulation) (Fig. 5D). In conclusion, PG1842 was upregulated in the ΔvimA mutant at stationary phase compared to that at exponential growth phase. PG1842 is the closest phylogenetic relative to VimA. The relatedness of VimA with the other putative acetyltransferases was evaluated. A phylogenetic tree of the six annotated acetyltransferases, namely, VimA, PG1088, PG1254, PG1358, PG1761, and PG1842, reconstructed with the neighbor-joining method, showed two related clusters (Fig. 6). Cluster 1 consisted of VimA, PG1842, and PG1254, while cluster 2 included the other three proteins, PG1358, PG1761, and PG1088. VimA showed the closest molecular relatedness to PG1842, followed by PG1254 (Fig. 6). jb.asm.org 6




FIG 4 Immunoblot analysis of the cell lysate (CL) and extracellular (ECL) protein fractions from P. gingivalis W83, ΔvimA, and ΔvimA-ΔPG1842 strains. CL and ECL fractions were collected from the bacteria grown to either exponential (OD60 of ⬃0.8) or stationary (OD600 of ⬃1.6) growth phase. Equivalent protein samples were separated on a 4 to 12% NuPAGE bis-Tris gradient gel and detected by immunoblotting with anti-RgpB (A) or anti-Kgp (B) antibodies. The positions of high-molecular-weight inactive proenzyme forms and active catalytic domain bands (shown by white arrows) are indicated. The inactive proenzyme species did not process into an active catalytic domain band in the ΔvimA-ΔPG1842 double mutant at stationary phase.


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FIG 5 Quantitative real-time PCR showing expression levels (fold change) of acetyltransferase genes PG1088, PG1254, PG1358, PG1761, and PG1842 in the W83 wild type (A) and ΔvimA mutant (B) at stationary phase compared to those at exponential phase and in the ΔvimA mutant compared to that in W83 at exponential phase (C) and at stationary phase (D). Total RNA from W83 and the ΔvimA mutant was extracted from either exponentially growing bacterial cells (OD600 of ⬃0.8) or stationary-phase cells (OD600 of ⬃1.6). Fold change in expression levels was calculated by the 2⫺ΔΔCT method. Gene expression levels were normalized to those of 16S rRNA transcripts. Data represent the averages from three independent experiments. Each experiment was done in triplicates. Error bars represent the standard deviations from the means.

Acetyltransferase domain of VimA is structurally similar to that of PG1842. The protein models for acetyltransferase domains of VimA, PG1088, PG1254, PG1358, PG1761, and PG1842 were predicted using I-TASSER. Furthermore, the template modeling (TM)-align algorithm was used to align the predicted domain structure of VimA with those of PG1088, PG1254, PG1358, PG1761, and PG1842. TM-align identifies the structural alignment between protein pairs on the basis of the TM score. The TM score measures the similarity between two protein structures and shows the difference as a score between 0 and 1, where 1 indicates an ideal match between two structures (33). Generally, scores below 0.20 indicate randomly chosen unrelated proteins, whereas structures with a score higher than 0.5 represent two proteins with approximately the same folds (33). As shown in Fig. 7, the acetyltransferase domain of VimA was most similar to those of PG1254 and PG1842, with TM scores of 0.63788 and 0.60489, respectively. Recombinant VimA and PG1842 acetylate pro-RgpB. If PG1842 is a functional VimA homolog and plays a role in gingipain activation, it should be possible for both VimA and PG1842 acetyltransferases to acetylate the gingipains. To examine the potential function of PG1842, both VimA and PG1842 proteins were overexpressed as C-terminal His tag fusion proteins in Escherichia coli. An in vitro acetyltransferase assay was performed with VimA/PG1842, pro-RgpB substrate, and an acetyl-CoA donor. Following the assay, tandem mass spectroscopy was used to identify acetylated residues in pro-RgpB. Acetylated peptides were identified by their mass increment of 42 Da compared to that of unacetylated peptides. As shown in Fig. 8, recombinant VimA and PG1842 acetylated Y230, K247, and K248 residues in a pro-RgpB substrate purified from the vimA-defective mutant FLL92 (Fig. 8A). A representative spectrum for tyrosine acetylation (Y230) is also shown (Fig. 8B). December 2018 Volume 200 Issue 24 e00385-18

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FIG 6 Phylogenetic tree of the putative GNAT family N-acetyltransferases in P. gingivalis W83 reconstructed with the neighbor-joining method by using the program MEGA, version 7.0. The optimal tree with the sum of branch lengths of 6.156 is shown. The tree is drawn to scale, with branch lengths (next to the branches) in the same units as those of the evolutionary distances used to infer the phylogenetic tree. The evolutionary distances were computed using the Poisson correction method and are in the units of the number of amino acid substitutions per site. VimA showed close molecular relatedness to PG1842.

DISCUSSION The regulation of gingipain activity in P. gingivalis occurs at multiple levels, including at the level of gingipain gene expression, the processing of an inactive proenzyme to an enzymatically active form, by secretion, and by glycosylation or another unknown modification of the proteins. Although the role of posttranslational processing during gingipain maturation and activation in P. gingivalis is well accepted, the underlying mechanism(s) of biogenesis is unclear. Only a few of the components involved at the posttranslational level are known. Additionally, as the secretion of these proteins in P. gingivalis involves a newly identified type IX secretion system (13), the complex mechanism of translocation through the outer membrane for secretion or attachment to the membrane is beginning to emerge. A recent report on the acetylome of P. gingivalis confirms that the three gingipain species, RgpA, RgpB, and Kgp, are acetylated, suggesting that acetylation may play role in gingipain biogenesis (27). This is consistent with our previous report which demonstrated that the vimA gene product, a putative acetyltransferase, modulates gingipain activity in P. gingivalis (29, 30). Although there was no gingipain activity in the ΔvimA mutant at exponential phase, we did see late-onset activity at stationary phase (Fig. 2). Because the VimA protein is a putative acetyltransferase, there is the possibility that a functional homolog that is upregulated/activated in the stationary growth phase plays a role in the late onset of gingipain activity in the ΔvimA mutant. In this report, we confirmed that PG1842 is a functional VimA homolog. Scanning of the P. gingivalis W83 genome identified five putative GNAT family proteins, including PG1088, PG1254, PG1358, PG1761, and PG1842 (Fig. 3A). To address the function of these proteins in the ΔvimA mutant, we deleted each of the genes in the ΔvimA background and then investigated if any of the double mutations affected the late onset of gingipain activity. Only one of the double isogenic mutants (the ΔvimAΔPG1842 mutant) showed the absence of late-onset gingipain activity at stationary December 2018 Volume 200 Issue 24 e00385-18

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FIG 7 Structural similarity between acetyltransferase domain of VimA with domains of PG1088, PG1254, PG1358, PG1761, and PG1842. The protein model for each domain was predicted by I-TASSER. The predicted model of VimA was aligned with models of PG1088, PG1254, PG1358, PG1761, and PG1842 using the TM-align server. The template modeling (TM) score for each protein pair is shown. On the basis of TM scores, the acetyltransferase domain of VimA is more closely related to PG1254 and PG1842.

phase (Fig. 2B), suggesting that PG1842 may be able to complement the functional role of VimA protein at stationary phase. Acetylation was originally discovered as a modification of histones in eukaryotes but recently appears as one of the commonly studied posttranslational modifications in prokaryotes (16). GNAT is a large family of enzymes typically catalyzing acetylation by using acetyl-CoA as the acetyl donor (16). This family is conserved and contains orthologous proteins from eukaryotes, prokaryotes, and archaea. The GNAT family of proteins has been classified into subfamilies on the basis of their substrate specificity. The primary sequences of these proteins show very low sequence identity, but they share a typical GNAT fold (16). These proteins appear to be versatile in terms of the substrates they target (such as regulatory proteins, small metabolites, and antibiotics) and the various functions they perform. Different studies indicate that subfamilies may use divergent regions flanking the conserved core (␣-␤-␤-␤-␣) of the GNAT fold to recognize a specific substrate(s) (34) and may use different chemical strategies to catalyze the acetylation reaction (34). In bacteria, GNAT enzymes affect many classes of proteins, including the enzymes of central metabolism, ribosomal proteins, cell wall homeostasis enzymes, and virulence-related proteins (16, 19, 24, 26). Our transcriptional analysis showed that except for PG1254, the other acetyltransferase genes (PG1088, PG1358, PG1761, and PG1842) were upregulated in the ΔvimA mutant at stationary phase compared to that at exponential growth phase (Fig. 5B). This raises the question as to why only PG1842 but not the other proteins compensates for VimA, given that all four genes were upregulated in the ΔvimA mutant at stationary phase. On the basis of their primary sequences, VimA is more distantly related to these proteins. In the GNAT family, PG1088 (Ac_7), PG1358 (Ac_3), PG1761 (Ac_1), and PG1842 (Ac_9) belong to different subfamilies (Fig. 3A) (32). It appears that these enzymes, including PG1842, have their own specific substrates, but due to some December 2018 Volume 200 Issue 24 e00385-18

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FIG 8 VimA and PG1842 acetylate pro-RgpB. (A) Summary of MS/MS identification of pro-RgpB. Acetylated (Ac) residues are indicated. The arrow indicates the cleavage site between R229 and Y230. (B) Representative spectra showing tyrosine acetylation by VimA and PG1842. (i) Tandem mass spectrum of the precursor ion at m/z 433.219 (MH2⫹) corresponding to the peptide sequence shown on the top right in the spectrum in which YTPVEEK is unmodified in the RgpB substrate. (ii) Tandem mass spectrum of the precursor ion at m/z 454.225 (MH2⫹) corresponding to the peptide sequence shown on the top right in the spectrum in which YTPVEEK is acetylated in the RgpB substrate reacted with VimA; typically, the immonium ion at m/z 178.087 confirms tyrosine acetylation in contrast to an unmodified tyrosine whose immonium ion is m/z 136.078. (iii) Tandem mass spectrum of the precursor ion at m/z 454.225 (MH2⫹) corresponding to the peptide sequence shown on the top right in the spectrum in which YTPVEEK is acetylated in the RgpB substrate reacted with PG1842. The measurement accuracy for precursor ions is smaller than 5 ppm and is smaller than 10 ppm for production ions.

similarity between PG1842 and VimA, PG1842 can also work on the VimA substrate RgpB. We hypothesize that PG1842 may fold into a tertiary structure similar to that of VimA that enables it to share the substrate for the acetylation reaction. Although this hypothesis remains to be tested experimentally, it is noteworthy that the tertiary December 2018 Volume 200 Issue 24 e00385-18

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structure of the acetyltransferase domain of VimA is most similar to those for PG1254 and PG1842 (Fig. 7). As PG1254 is downregulated in the ΔvimA mutant at stationary phase (Fig. 5B), PG1842, with a similar tertiary structure, may work on VimA’s substrate. Proteins from a common ancestor may share similar functions. It is possible that VimA and PG1842 have evolved from the same ancestor and later diverged to act on different substrates. In fact, we have shown that phylogenetically, VimA is most closely related to PG1842 (Fig. 6). One intriguing observation is that PG1842 is also upregulated (though at low levels) at exponential phase in the ΔvimA mutant (Fig. 5C). PG1842 RNA is transcribed at both exponential and stationary phases, but it is likely that the PG1842 protein is activated to work on gingipains only at the stationary growth phase. This raises the question of a likely mechanism for the activation of PG1842 at stationary phase. Our preliminary studies in the laboratory have shown that certain residues in the recombinant PG1842 protein can be acetylated; however, in contrast to that for VimA, there is no autoacetylation in the presence of acetyl-CoA as the acetyl donor (H. M. Fletcher, unpublished data). This might indicate that PG1842 needs to be acetylated to be active as an acetyltransferase and to act on other substrates. Reports have shown that acetylated proteins are more abundant in bacteria at stationary growth phase, and after the resuspension of such cells in fresh medium, the numbers of acetylated proteins are reduced (17). The acetylation of the DNA repair protein Ku in Mycobacterium smegmatis increases from exponential to stationary growth phase (35). In Bacillus subtilis, the rod shape-determining protein MreB is acetylated at three acetylation sites, with K240 acetylation significantly increased when the bacteria enters into stationary phase (36). Wild-type cells get shorter and narrower in stationary phase and also in exponential phase in the MreB K240Q (Q mimics the acetylated K residue) mutant. During stationary phase, the K240R mutant (R mimics K in unacetylated form) cells are similar to exponential-phase cells, perhaps due to the inability to acetylate the MreB K240R protein (36). On the basis of the above observations, it is possible that PG1842 gets activated by acetylation only at stationary phase. Therefore, although the protein is synthesized during both exponential and stationary phases, it may act on gingipains only during the stationary phase. To confirm this, we need to compare the acetylation profiles of PG1842 in vivo at both exponential and stationary growth phases. More experiments are needed to support this hypothesis. Mass spectroscopy data showed that both VimA and PG1842 are able to acetylate pro-RgpB at residues Y230 (O-acetylation), K247, and K248 (N␧-acetylation). Acetylation consists of the transfer of an acetyl group either at the proteins’ N terminus (N␣acetylation) or on the side chains of the lysine (N␧-acetylation), serine, and threonine (O-acetylation) residues (37). N␧-Acetylation of lysine residues is well characterized, whereas N␣-acetylation and O-acetylation have not been extensively studied in bacteria. O-acetylation by YopJ family proteins plays an important role in virulence and promotes bacterial survival in host organisms. In Yersinia pestis, the type III secretion system directly injects YopJ effector protein into the immune cells of the infected host. YopJ acetylates the active site serine and threonine residues of host mitogen-activated protein kinases. This acetylation blocks phosphorylation and therefore prevents the signaling pathway in the host cell (38, 39). To the best of our knowledge, O-acetylated proteins have not been characterized in bacteria. This is mainly due to the unstable nature of the O-N-acyl transfer reaction, which prevents its detection (40). With the development of exceptionally sensitive technologies for detecting protein modifications, O-acetylation is far easier to identify. Recently, a quantitative acetylome analysis in Mycobacterium tuberculosis identified 2,349 O-acetylation and 141 N␧-acetylation sites, derived from 953 unique proteins (41). Thus, M. tuberculosis O-acetylation was significantly more abundant than N␧-acetylation. This fact may support the notion that O-acetylation, including that of tyrosine, is involved in regulating more multiple biological functions than expected, but a strong chemical instability of O-acetylation makes its detection difficult compared to that for N-acetylation (40). How one acetyltransferase enzyme is catalyzing N-acetylation (K247 and K248) and O-acetylation jb.asm.org 12




(Y230) reactions on one substrate is not clear. Earlier published reports showed that YopJ (Ac_14; pfam03421) of Y. pestis can acetylate the serine, threonine, and lysine residues of eukaryotic mitogen-activated protein kinases (38). There are apparently no reports of tyrosine acetylation in bacteria. The significance of Y230 acetylation is not clear in the case of RgpB activation/maturation. Residue Y230 is of particular interest in RgpB. The activation of RgpB requires autolytic processing. Mikolajczyk et al. (11) previously showed that full in vitro activation of the recombinant RgpB involved three sequential autolytic processing steps. The first two steps comprise the removal of the N-terminal propeptide sequentially, followed by the removal of the CTD at the third step. During the first two steps, the N-terminal prodomain is removed by cleaving between residues R229 and Y230 (denoted by arrow in Fig. 8A). Since the VimA and PG1842 acetyltransferases are predicted to be localized in the cytoplasm (https://www.psort.org), we speculate that pro-RgpB gets acetylated at Y230 in the cytoplasm. It is possible that the acetylation of Y230 affects the cleavage at R229. A defect in N-terminal processing would likely affect the subsequent removal of the CTD, thus resulting in the secretion of the acetylated proenzyme species into the outer environment. The removal of the acetyl group at Y230 by an unknown deacetylase will facilitate the N-terminal processing of the prodomain and subsequent removal of the CTD, which can result in the attachment of the mature gingipains to the outer membrane. The likely deacetylase playing a role in gingipain biogenesis might involve the hypothetical protein encoded by vimE, which is part of the vimA transcriptional unit (28). VimE is a member of the CE4 superfamily and is predicted to be a NodB-like deacetylase. VimE has been shown to play role in gingipain biogenesis (42). A more detailed mechanism is being further investigated in our lab. In summary, our data confirm that VimA and PG1842 are acetyltransferases. The data also provide evidence that acetylation plays a role in the gingipain activation/maturation pathway. Work is under way to determine the importance of each acetylated residue, including Y230. This will expand the understanding regarding the nature of acetylated proteins in P. gingivalis and open a new avenue of research for exploring the role of protein acetylation in gingipain biogenesis. MATERIALS AND METHODS Bacterial strains, plasmids, and growth conditions. The bacterial strains and plasmids used in this study are listed in Table 1. All P. gingivalis strains were grown in brain heart infusion (BHI) broth supplemented with yeast extract (0.5%), hemin (5 ␮g/ml), vitamin K (0.5 ␮g/ml), and DL-cysteine (0.1%; Sigma). Defibrinated sheep blood (5%) and agar (1.5%) were used in blood agar plates. Escherichia coli strains were grown in Luria-Bertani (LB) broth. P. gingivalis strains were maintained in an anaerobic chamber (Coy Manufacturing) in 10% H2, 10% CO2, and 80% N2 at 37°C. Growth rates for P. gingivalis and E. coli strains were determined spectrophotometrically by measuring the optical density at 600 nm (OD600). When needed, antibiotics were added at the following concentrations: 10 ␮g/ml erythromycin and 3 ␮g/ml tetracycline for P. gingivalis, and 100 ␮g/ml ampicillin for E. coli. Construction of bacterial strains. An isogenic vimA mutant in P. gingivalis W83 was constructed by using a long-PCR-based fusion method as previously described elsewhere (43). Briefly, fragments containing 498 bp upstream and 507 bp downstream of vimA were PCR amplified from W83 chromosomal DNA. The promoterless erythromycin resistance cassette (ermF) without a transcriptional terminator was amplified from the plasmid pVA2198 (44) with primers containing overlapping nucleotides for the upstream and downstream vimA fragments. The upstream fragment, ermF, and downstream fragments were fused together, and the purified fusion product was electroporated into W83 as previously described (29). The cells were plated on BHI agar plates containing 10 ␮g/ml of erythromycin and incubated at 37°C for 7 days. The correct gene replacement in the erythromycin-resistant mutant was confirmed by PCR and DNA sequencing. To create the double mutants in the ΔvimA background, ⬃500-bp upstream and downstream fragments of the respected genes were fused to the promoterless tetracycline resistance cassette (tetQ) and then introduced into P. gingivalis FLL102 (ΔvimA::ermF) by electroporation (43). The mutants were selected on BHI agar plates containing 10 ␮g/ml of erythromycin and 3 ␮g/ml of tetracycline. The correct gene replacement in the erythromycin- and tetracycline-resistant mutant was confirmed by PCR and DNA sequencing. The primers used in this study are listed in Table S1 in the supplemental material. Gingipain activity assay. The presence of Arg-X- and Lys-X-specific cysteine protease activity (Rgp and Kgp, respectively) was determined by using 1 mM BAPNA (N␣-benzoyl-DL-arginine-p-nitroanilide) and 1 mM ALNA (Ac-Lys-p-nitroanilide HCl) as the substrates in an activated protease buffer (0.2 M Tris-HCl, 0.1 M NaCl, 5 mM CaCl2, 10 mM L-cysteine [pH 7.6]) (29). Substrates were individually added to P. gingivalis exponential-phase (OD600 of ⬃0.8) and stationary-phase (OD600 of ⬃1.6) culture samples, and December 2018 Volume 200 Issue 24 e00385-18

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TABLE 1 Bacterial strains and plasmids used in this study Strain or plasmid P. gingivalis strains W83 FLL92 FLL102 FLL122 FLL123 FLL124

FLL126

E. coli strains Top 10 BL21 Star(DE3)

Plasmids pT-COW pVA2198 pET102-TOPO pET102-vimA pET102-PG1842

Reference or source

Wild-type strain vimA::ermF-ermAM; an isogenic derivative of W83 ΔvimA::ermF; an isogenic derivative of W83 ΔvimA::ermF-ΔPG1088::tetQ; an isogenic derivative of FLL102 ΔvimA::ermF-ΔPG1254::tetQ; an isogenic derivative of FLL102 ΔvimA::ermF-ΔPG1358::tetQ; an isogenic derivative of FLL102 ΔvimA::ermF-ΔPG1761::tetQ; an isogenic derivative of FLL102 ΔvimA::ermF-ΔPG1842::tetQ; an isogenic derivative of FLL102

29 29 This study This study

Used for general cloning purpose F⫺ ompT hsdSB(rB⫺ mB⫺) gal dcm rne131 (DE3), used as a protein expression strain

Invitrogen Invitrogen

Apr tetQ Spr ermF-ermAM Apr, C-terminal His tag Apr, pET102 derivative expressing vimA Apr, pET102 derivative expressing PG1842

52 44 Invitrogen This study This study

This study This study This study


FLL125

Genotype and/or description

This study

the endpoint OD was measured at 405 nm against a blank sample containing no bacteria by using a microplate reader (Bio-Rad). P. gingivalis cell fractionation and immunoblot analysis. Total cell lysate and extracellular fractions were prepared from wild-type P. gingivalis W83 and its isogenic mutants according to a previously published protocol (42) with some modifications. Briefly, the cells were grown to exponential log phase (OD600 of ⬃0.8) and/or stationary phase (OD600 of ⬃1.6) and centrifuged at 10,000 ⫻ g for 1 h at 4°C. The proteins from the cell-free supernatants were precipitated with 80% ammonium sulfate. The ammonium sulfate protein precipitate was resuspended in 10 mM Tris-HCl (pH 7.4) and dialyzed extensively against the same buffer to remove ammonium sulfate (extracellular fraction). The cell pellets were washed in 10 mM Tris-HCl buffer (pH 7.4) and lysed with a French pressure cell press (American Instrument Company) in the presence of protease inhibitor. After centrifuging, the supernatants were designated the total cell lysates. The proteins from the cell lysates and extracellular fractions were separated on 4 to 12% gradient gels (Invitrogen) and transferred to Bio-trace nitrocellulose membranes. The blots were probed with antibodies against specific protease domains (anti-Kgp, anti-RgpA, and anti-RgpB) and species-specific secondary antibodies conjugated to horseradish peroxidase (Zymed Laboratories). Immunoreactive protein bands were detected according to the procedure described in the Western Lightning chemiluminescence reagent plus kit (Perkin-Elmer Life Sciences). Purification of the RgpB gingipain. The RgpB gingipain was purified according to the method described by Potempa and Nguyen with some modifications (45). Instead of acetone precipitation, 80% ammonium sulfate was used to precipitate the gingipain from the culture supernatant (extracellular fraction) of P. gingivalis FLL92 (vimA-defective mutant) grown to exponential phase (OD600 of ⬃0.8). In addition, four columns were used in the following order: Hi Load 16/60 Superdex 200, DEAE-cellulose anion-exchange column, arginine-Sepharose column, and a Superdex 200HR 10/30 column. RNA isolation and qRT-PCR. Log-phase cultures (OD600 of ⬃0.8) of different P. gingivalis strains were used to isolate total RNA with the SV Total RNA isolation kit (Promega) according to the manufacturer’s instructions. DNA contamination in the RNA preparation was cleaned by treating the RNA with DNase (Qiagen). The DNase-treated RNA was checked for DNA contamination by PCR, and the integrity of the RNA was analyzed visually by gel electrophoresis. RNA concentrations and purity were measured with a Nanodrop 2000 (Fisher Scientific). cDNA was synthesized using a Transcriptor High Fidelity cDNA synthesis kit (Roche Corp.) according to the manufacturer’s protocol. qRT-PCR was performed with the SYBR green kit (Life Technologies), and real-time fluorescence was detected by the Applied Biotechnology QuantStudio 7 Flex real-time PCR system (Fisher Scientific) according to the manufacturer’s instructions. Forty cycles of the following program were used: 50°C for 2 min, 95°C for 5 min, 95°C for 15 s, 54°C for 30 s, and 72°C for 30 s. The melting curve profiles were reviewed to verify single peaks for individual samples. Each experiment was done in triplicates, and experiments were independently performed three times with comparable results. The 16S rRNA gene was used as a reference to normalize gene expression. The 2⫺ΔΔCT method was used (46) to evaluate the relative gene expression in different strains and/or December 2018 Volume 200 Issue 24 e00385-18

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different growing phases (exponential versus stationary). The primers used for qRT-PCR (Table S1) were designed using the PrimerQuest tool from Integrated DNA Technologies (IDT). Bioinformatics analysis. The phylogenetic relationship between annotated acetyltransferases PG1088, PG1254, PG1358, PG1761, PG1842, and VimA was analyzed by using the MEGA program, version 7.0 (47). The evolutionary history was derived by using the neighbor-joining method (48). The evolutionary distances were computed with the Poisson correction method (49) and are in the units of the number of amino acid substitutions per site. All positions containing gaps and missing data were eliminated during the analysis of the six protein sequences. There were a total of 116 positions in the final data set. The protein models were predicted by the I-TASSER server (https://zhanglab.ccmb.med.umich.edu/ I-TASSER/) (50), whereas the TM-align server (https://zhanglab.ccmb.med.umich.edu/TM-align/) (51) was used for comparison studies of two different models. The protein domains were predicted by the NCBI CDD database (32). Cloning, expression, and purification of recombinant VimA and PG1842 proteins. P. gingivalis genes, including vimA and PG1842, were PCR amplified from W83 genomic DNA and cloned into expression vector pET102/D-TOPO by using a Champion pET102 Directional TOPO expression kit (Life Technologies). The DNA sequences of vimA and PG1842 in the recombinant plasmids were confirmed by sequencing. The recombinant plasmids were then transformed into E. coli BL21 Star(DE3) cells for expression. E. coli BL21 cells harboring the recombinant plasmids were grown at 37°C in LB medium supplemented with 100 ␮g/ml of ampicillin until the culture reached mid-log phase (OD600 of ⬃0.4). The cells were harvested by centrifugation 4 h after induction (with 1 mM IPTG [isopropyl-␤-Dthiogalactopyranoside]) and stored at ⫺80°C. The insoluble recombinant proteins were purified from the pellet fraction using nickel-nitrilotriacetic acid (Ni-NTA) beads according to the manufacturer’s instructions (Qiagen). To renature VimA, the eluted fractions were dialyzed against 4 M urea buffer (0.5 mM NaCl, 20 mM NaH2PO4, 4 M urea), followed by 2 M urea buffer (0.5 mM NaCl, 20 mM NaH2PO4, 2 M urea, and 0.5 M L-arginine) and 1 M urea buffer (0.5 mM NaCl, 20 mM NaH2PO4, 1 M urea, and 0.5 M L-arginine). Finally, the protein was dialyzed twice in the same buffer without any urea. PG1842 protein was renatured by dialyzing the eluted protein fractions against 4 M urea buffer (50 mM Tris-HCl, 150 mM NaCl, 4 M urea, pH 8.0), followed by 2 M urea buffer (50 mM Tris-HCl, 150 mM NaCl, 2 M urea, 0.5 M L-arginine, pH 8.0) and 1 M urea buffer (50 mM Tris-HCl, 150 mM NaCl, 1 M urea, 0.5 M L-arginine, pH 8.0). Finally, the protein was dialyzed twice in the same buffer without any urea. During dialysis, the lack of a visible precipitate indicated that the proteins were refolded properly. After dialysis, the recombinant proteins were run on SDS-PAGE gels, concentrated via an Amicon Ultra centrifugal filter (Millipore) to the desired concentrations, mixed with 10% glycerol, aliquoted, and stored at ⫺80°C for further use. In vitro acetylation assay. In vitro acetylation assays were performed in a 30-␮l total reaction volume containing 50 mM Tris-HCl (pH 7.5), 500 ␮M acetyl-CoA (Sigma), 2 mM TLCK (Sigma), 7.5 ␮M recombinant VimA or PG1842 (as required) and 40 ␮M pro-RgpB as the substrate. TLCK (N␣-p-tosyl-Llysine chloromethyl ketone) was used to inhibit the residual activity of catalytic RgpB present in purified preparations of pro-RgpB. After incubating for 1 h at 37°C, the reactions were stopped by the addition of 4⫻ SDS sample buffer (Invitrogen). Mass spectrometry analysis. After in vitro acetylation, the proteins were separated by SDS-PAGE. The excised SDS-PAGE gel bands with proteins of interest were incubated with 50% methanol in distilled water to remove Coomassie blue dye, followed by two washes in distilled water. The gel bands were dried with Kimwipes and crushed into fine particles with a 100-␮l pipet tip. The crushed gels were immersed in 100 ␮l of 50 mM NH4HCO3 before 1 ␮g of trypsin (Sigma) was added for digestion at 37°C overnight. The digested peptides were extracted from the gels twice with 500 ␮l of acetonitrile. The extracts were dried via a reduced vacuum in a Speedvac and redissolved in 20 ␮l of 1% formic acid before being transferred to high-pressure liquid chromatography (HPLC) autosampler vials for analysis. Liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis was carried out on the Thermo hybrid linear ion trap-Orbitrap mass spectrometer (Orbitrap Elite; Thermo Scientific). Peptides were separated by online reverse-phase liquid chromatography (RPLC) by using a two-column setup: a trap column (2-cm long, C18, 100-␮m inside diameter [i.d.]; Thermo Scientific) followed by C18 analytical columns (ProteoPep II, 15-cm long, 75-␮m i.d., 15-␮m tip, 5-␮m particle size; Thermo Scientific) with a 400-nl/min flow rate and 165-min gradient (solvent A, 0.1% formic acid in water; solvent B, 0.1% formic acid in acetonitrile [Thermo Scientific]) from 2% to 95% solvent B. The Orbitrap mass analyzer was set to acquire data at a 60,000 resolution (full width at half maximum [FWHM]) for the parent full-scan mass spectrum (scan range, m/z 350 to 1,600) followed by high-energy collision dissociation (HCD) MS/MS spectra for the top 15 most abundant ions acquired at a 15,000 resolution. MS data analysis. Proteins were identified by PEAK 8.5 (Bioinformatics Solutions Inc., Waterloo, ON, Canada) to perform a de novo sequencing-assisted database search against customer provided database constructed from P. gingivalis protein PG_0882 (WP_005874177.1), PG_0506 (WP_010956050.1), and PG_1842 (WP_043876453.1). The acetylation of lysine (K), serine (S), threonine (T), and tyrosine (Y), the oxidation of methionine, and the deamination of asparagine (D) and glutamine (Q) were set as variable modifications. Trypsin was the selected protease, and up to two missed cleavages were used in the analysis. Mass tolerance for the precursor ions was 5 ppm and was 10 ppm for the MS/MS. Acetylated peptides were further manually confirmed by comparing their detected fragmentation ions with theoretic counterparts and guaranteeing the appearance of m/z 126.091, 102.055, 116.071, and 178.087 corresponding to acetylated K, S, T, and Y. jb.asm.org 15

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SUPPLEMENTAL MATERIAL Supplemental material for this article may be found at https://doi.org/10.1128/JB .00385-18. SUPPLEMENTAL FILE 1, PDF file, 0.3 MB. ACKNOWLEDGMENTS We thank the members of our lab for their critical inputs. This work was supported by grants R-56-DE13664, DE019730, DE022508, and DE022724 from the NIDCR (to H.M.F.). All authors declare no conflict of interest for this work.

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