Bacterial Infections

The Porphyromonas gingivalis Hybrid Cluster Protein Hcp Is Required for Growth with Nitrite and Survival with Host Cells

B. Ross Belvin, Qin Gui, Justin A. Hutcherson, Janina P. Lewis
Marvin Whiteley, Editor
DOI: 10.1128/IAI.00572-18

ABSTRACT

Although the periodontal pathogen Porphyromonas gingivalis must withstand high levels of nitrosative stress while in the oral cavity, the mechanisms of nitrosative stress defense are not well understood in this organism. Previously we showed that the transcriptional regulator HcpR plays a significant role in defense, and here we further defined its regulon. Our study shows that hcp (PG0893), a putative nitric oxide (NO) reductase, is the only gene significantly upregulated in response to nitrite (NO2) and that this regulation is dependent on HcpR. An isogenic mutant deficient in hcp is not able to grow with 200 μM nitrite, demonstrating that the sensitivity of the HcpR mutant is mediated through Hcp. We further define the molecular mechanisms of HcpR interaction with the hcp promoter through mutational analysis of the inverted repeat present within the promoter. Although other putative nitrosative stress protection mechanisms present on the nrfAH operon are also found in the P. gingivalis genome, we show that their gene products play no role in growth of the bacterium with nitrite. As growth of the hcp-deficient strain was also significantly diminished in the presence of a nitric oxide-producing compound, S-nitrosoglutathione (GSNO), Hcp appears to be the primary means by which P. gingivalis responds to NO2−-based stress. Finally, we show that Hcp is required for survival with host cells but that loss of Hcp has no effect on association and entry of P. gingivalis into human oral keratinocytes.

INTRODUCTION

Porphyromonas gingivalis is a Gram-negative anaerobic bacterium implicated in the development of periodontal diseases (1, 2). P. gingivalis utilizes its many virulence factors to deregulate the host immune response and upset host-microbiome homeostasis, thereby creating a proinflammatory environment characteristic of periodontal diseases. Understanding the capacity of P. gingivalis to persist in the oral cavity is vital to better understanding the progression of periodontal disease and may aid in development of therapeutic measures to reduce levels of the bacterium and prevent periodontitis. The oral cavity is rich in reactive nitrogen species (RNS). The primary sources of RNS are derived from the host and diet and can take the form of species such as nitrate (NO3−), nitrite (NO2−), and nitric oxide (NO) (3, 4). After a meal, bacteria in the oral cavity can produce nitrogen species as metabolic by-products (5). Members of the host oral microbiome are capable of reducing diet-derived nitrate to nitrite through the use of nitrate reductase systems, creating an environment with elevated nitrite concentrations that can exceed 1 mM after a nitrate-rich meal (4, 6, 7). Increased intake of nitrate also leads to elevated levels of nitrite in blood and serum (7). In the oral cavity, heightened levels of nitrite may lead to the chemical generation of NO through the acidification of nitrite (3). Another source of NO in the oral cavity is host cells, which produce NO- and O2−-based species in response to microbial infection. This is part of the innate immune response and is regulated by host nitric oxide synthases (NOS) (8). P. gingivalis has been shown to promote NO generation via the activation of inducible NOS (iNOS) in both immune and nonimmune cells (9–11).

Despite the high concentrations of RNS in the oral cavity, the mechanisms P. gingivalis uses to persist in this environment are not completely understood. We have previously shown that the HcpR mutant is not able to grow with 1 mM nitrite, a concentration normally encountered in the oral cavity (12). In addition, the mutant lost its ability to survive with host cells. These data were further verified via a transposon insertion library which also identified an HcpR mutant as having reduced ability to colonize host cells as well as survive in mice, indicating that the regulator plays an important role in virulence of the bacterium and survival in the host (13). Although we have shown that this regulator was indispensable for upregulation of the redox enzyme known as the hybrid cluster protein (Hcp) (also known as prismane protein), the entire regulon of HcpR, as well as the role of Hcp, has not been defined; thus, the exact mechanism of the HcpR sensitivity to nitrosative stress remained unknown. Study in the Fletcher laboratory has shown that Hcp is required for P. gingivalis growth with NO; however, its exact role in the nitrosative stress response is not known (14).

Hcp has been found in an array of Gram-negative obligate anaerobes (Bacteroides and Desulfovibrio spp.) and facultative anaerobes (Escherichia coli and Pseudomonas spp.), where it is induced by NO3−, NO2−, or NO (15–17). It has been implicated to act as a putative hydroxylamine reductase (14, 18). Other studies suggest that Hcp is a high-affinity NO reductase in enteric bacteria that protects cytoplasmic proteins and DNA from nitrosative stress damage (16). Recent studies in E. coli also suggest that Hcp plays an important role as a regulator of endogenous protein nitrosylation, thereby regulating nitrogen-based metabolism and the bacterial response to reactive by-products produced during nitrogen metabolism (19).

Besides carrying hcp, the genome of P. gingivalis codes for a number of putative proteins that are hypothesized to be involved in nitrosative stress resistance. One such operon, composed of PG1820 and PG1821, codes for a putative NrfAH nitrite reductase system. This is a large periplasmic complex that utilizes nitrite as an electron sink, reducing it to ammonia to create a membrane potential (20–22). The nrf operon has been shown to play a role in protection against nitrosative stress in other Gram-negative anaerobic and microaerophilic bacteria (23–25). However, its function and role in nitrosative stress defense in P. gingivalis have not been investigated.

In this study, we further investigated the mechanisms involved in the response of P. gingivalis to nitrosative stress. First, we ascertain changes in gene expression affected by the loss of HcpR. A transcriptome sequencing (RNA-seq) approach was utilized to better characterize the bacteria’s response to nitrite and better understand HcpR’s role in the nitrosative stress response. We verified the regulation of hcp by HcpR and demonstrated the biological role of Hcp in nitrosative stress protection. We show that Hcp is essential for the growth of P. gingivalis in physiological concentrations of nitrite and that it is also vital for its survival with host cells. Furthermore, we verify that HcpR is a direct regulator of hcp in response to nitrosative stress. Genes that are hypothesized to play a role in nitrosative stress protection, nrfAH, were also investigated and were compared to the hcp knockout strain. This work further sheds light on the nitrosative stress response in P. gingivalis and helps fill the gap in our understanding of its mechanisms.

RESULTS

Transcriptional response of P. gingivalis to nitrite.To better understand the bacterial response to nitrite and to identify putative members of the HcpR regulon, a whole-transcriptome sequencing approach was taken. Libraries of the W83 and V2807 (hcpR deletion mutant) strains with and without exposure to 200 μM nitrite were generated and then sequenced. The sequencing results were then compared, and the top 20 most upregulated and downregulated genes are shown in Tables 1 to 4.

View this table:
TABLE 1

Genes upregulated and downregulated by nitrite in P. gingivalis W83

View this table:
TABLE 2

Genes upregulated and downregulated by nitrite in P. gingivalis V2807

View this table:
TABLE 3

Differential expression of genes in the W83 and V2807 strains in the absence of nitrite

View this table:
TABLE 4

Differential expression of genes in P. gingivalis W83 and V2807 strains in the presence of 200 μM nitrite

Table 1 lists differential expression of genes with or without nitrite in wild-type P. gingivalis W83. The most upregulated gene, by a large margin, is hcp (PG0893), at 144.15-fold higher in the nitrite-treated sample, which agrees with quantitative reverse transcription-PCR (qRT-PCR) studies (Table 5; also see Fig. S1 in the supplemental material). The second most upregulated gene is just upstream of hcp, PG0890, and it is expression is increased by only 2.91-fold. This gene codes for a putative RadC protein involved in DNA repair. However, the increase of PG0890 may be artificial due to its proximity to the very highly upregulated PG0893 (hcp). PG1979 expression is increased 2.59-fold. This gene codes for a small hypothetical protein with unknown function that is found only in P. gingivalis. After these 3 genes, there are no other genes that are upregulated more than 2-fold. The presence of nitrite does not exert any change in the expression of hcpR at the transcriptional level.

Among the downregulated genes in the wild-type strain, no gene is more than 2-fold downregulated, and no single gene has a significant P value (P < 0.05). PG1534 is the gene that is most downregulated in W83 when exposed to nitrite; however, it is only regulated 1.67-fold. Nitrite appears to have very little effect on the downregulation of genes at the transcriptional level.

Table 2 lists differential expression of genes with or without nitrite in the V2807 strain (hcpR mutant). Two of the significantly upregulated genes are both subunits of the cytochrome d ubiquinol oxidase, cydA and cydB. These genes are upregulated comparably at 3.61- and 3.23-fold, respectively. Both the cydA and cydB genes are regulated and expressed as a transcriptional unit, and this is reflected in their similar fold changes. PG1222 codes for a hypothetical protein of unknown function that is well conserved among P. gingivalis strains. Of note, upregulation of hcp is not present without HcpR.

The most downregulated gene in the hcpR-deficient V2807 strain is the RpmH ribosomal protein at 2.6-fold; however, this does not have a significant P value. Of the most downregulated genes, PG0505, PG1904, and PG1484 are the only ones that have a significant P value. Unfortunately, all three of these genes code for hypothetical proteins of unknown function.

The expression profiles of the wild-type V2802 and hcpR-deficient V2807 strains are compared in Table 3. Although there are a number of genes that have >2.0-fold changes in transcript levels, there were no gene transcripts that significantly differed (P < 0.05) between the wild type and mutant. PG1555, a conserved membrane protein of unknown function, is the most upregulated gene at 3.31-fold. PG1570 codes for a rhodanese-like protein that is hypothesized to play a role in cyanide detoxification.

Finally, the transcriptomes of the W83 and V2807 strains after exposure to nitrite are compared in Table 4. hcp gene (PG0893) expression is 250-fold higher in the wild-type strain than in the V2807 strain. This gene is by far the most differentially regulated. The second most differentially regulated gene, PG1556, is expressed at a 5.43-fold higher level in the wild type than in the mutant. This gene is a hypothetical protein that is conserved in P. gingivalis and contains a well-conserved domain of unknown function. PG0524 expression is 4.39-fold higher in the wild-type strain and codes for a conserved hypothetical protein found in P. gingivalis with no known function. Genes that appear to have higher expression in the mutant when exposed to nitrite again include the cydA and cydB genes.

The levels of hcp transcript were confirmed by qRT-PCR (Table 5). After a 15-min exposure to nitrite, there is a 250-fold increase in hcp transcript levels in the W83 strain. This upregulation is absent from the V2807 strain. Furthermore, we also observed an increase in Hcp at the protein level when a FLAG-tagged hcp complemented strain was exposed to nitrite (Fig. S1A). Hydroxylamine did not stimulate the expression of hcp. These data suggest that Hcp and HcpR play a central and irreplaceable role in the bacterial response to nitrite-based stress.

View this table:
TABLE 5

Differential expression of P. gingivalis hcp in the presence of nitrosative stress

Sensitivity of the hcp mutant strain of P. gingivalis to nitrosative stress.Previously it was shown that the V2807 hcpR-deficient strain of P. gingivalis was susceptible to physiological concentrations of nitrite. We compared the abilities of the wild-type, V3239, V3200, and V3205 strains to grow in nitrite (Fig. 1). Concentrations as low as 0.2 mM NO2− are capable of inhibiting the growth of the hcp mutant strain, and growth is completely abolished at 0.5 mM nitrite. In contrast to this, wild-type P. gingivalis is capable of growing at 2 mM nitrite with little to no inhibition of growth. This indicates that the hcp mutant strain is very sensitive to nitrite at levels that are typically found in the oral cavity. The V3200 and V3205 strains show significantly decreased growth in the samples with and without nitrite. Although the nrfAH mutants may be physiologically compromised, the decreased rate of growth is not dependent on nitrite. This implies that the primary driver behind the P. gingivalis response to nitrite is dependent on Hcp and HcpR and that under these conditions the nrfAH operon does not play a significant role in protection from nitrite-based stress.

FIG 1
FIG 1

Hcp is required for P. gingivalis growth with nitrite. Bacterial strains were grown anaerobically in mycoplasma medium for 24 h. All cultures were supplemented with appropriate concentrations of NaNO2−. Unsupplemented cultures served as controls. All graphs are averages from at least 3 biological replicates. (A) Wild-type P. gingivalis W83 was grown in 0, 1, 4, and 8 mM concentrations of NO2− to establish a baseline of nitrite sensitivity. (B) The V3239 hcp knockout strain of P. gingivalis was grown in 0, 0.2, 0.5, and 2.0 mM NO2−. There is no statistical significance in the growth at 0 mM nitrite between the wild type and the mutant. (C) The growth rates of the wild-type parental strain, nrfA knockout mutant (V3205), and nrfH knockout mutant (V3200) were compared in mycoplasma medium. (D) The growth rates of the wild-type parental strain, nrfA knockout mutant (V3205), and nrfH knockout mutant (V3200) were compared in the presence of 2 mM NO2−. For all studies, statistical significance was measured after 24 h (*, P < 0.05; **, P < 0.01). Abs, absorbance.

To determine whether the protective effects of Hcp toward nitrite occur intracellularly or extracellularly, we performed a dot blot analysis using a FLAG-tagged Hcp complemented strain. After exposure to nitrite, there was a noticeable increase in the intracellular fraction; however, Hcp was undetectable in the extracellular fraction (Fig. S2A). Furthermore, conditioned media derived from wild-type P. gingivalis grown with nitrite provided no protective effect to the hcp mutant strain (Fig. S2B). This indicates that the protective effect provided by Hcp toward nitrite stress occurs inside the cell and that Hcp is not secreted.

We also tested the ability of the hcp-deficient strain (V3239) to grow in the presence of NO-generating S-nitrosoglutathione (GSNO) species (Fig. 2). NO is a robust stimulator of hcp expression: nanomolar concentrations of NO are capable of eliciting a substantial increase in the upregulation of hcp at the transcript level, and this expression is dependent on HcpR (Table 5 and Fig. S1). This implies that hcp plays an important role in the cellular defense against NO. Previous studies have used NONOate supplemented over 1 h to test susceptibility of the hcp knockout strain to growth in NO, revealing that hcp is important in NO stress resistance. GSNO is a more stable NO donor and has a much lower decomposition rate than NONOate, and it mimics a low exposure over time rather than a burst of NO (26, 27). As shown in Fig. 2A, at 25 μM the wild-type strain grows at a slightly reduced rate at early time points; however, the culture continues to grow over the course of 24 h. At 25 μM the hcp-deficient strain shows little to no growth at the 3- and 6-h time points, but as the levels of NO decrease over the course of 24 h, it is capable of growing to reduced levels compared to those of the wild-type parental strain. At 100 μM the W83 and V2807 strains exhibit little to no growth at the 3- and 6-h time points and greatly reduced growth after 24 h.

FIG 2
FIG 2

Hcp is required for P. gingivalis growth with GSNO. All cultures were grown anaerobically in mycoplasma medium for 24 h. All experiments are representative of 3 biological replicates. (A) Wild-type and hcp null (V3239) strains of P. gingivalis grown in 25 μM GSNO (*, P < 0.05). (B) Wild-type and hcp null (V3239) strains of P. gingivalis grown in 100 μM GSNO.

Binding of HcpR to the hcp promoter.To gain insights into the transcriptional regulation of hcp, an in vivo chromatin immunoprecipitation (ChIP) assay was performed. It was previously posited that HcpR is the primary and direct transcriptional regulator of hcp in response to nitrite (12). This is confirmed through a FLAG tag-based ChIP assay using tagged HcpR expressed in the V3239-deficient strain. After exposure to nitrite and immunoprecipitation, PCR was utilized to probe for the hcp promoter region (Fig. 3A). The promoter region was successfully pulled down and enriched in the reaction mixtures using full-length HcpR (lane 2); however, it was not found in the negative control using HcpR lacking the putative DNA binding domain (lane 3).

FIG 3
FIG 3

HcpR binds to the promoter region of hcp. (A) ChIP assay. Recombinant Flag-tagged HcpR was expressed in the hcpR null strain and immunoprecipitated. PCR was used to confirm coimmunoprecipitation of the hcp promoter (prom.). A truncated form of HcpR lacking the DNA binding domain was utilized as a negative control. Experiments are representative of 3 biological replicates. Lane 1, ladder; lane 2, wild-type HcpR ChIP; lane 3, L156* HcpR ChIP; lane 4, genomic DNA PCR. (B) The hcp-deficient strain was complemented using the Pg108 vector with the hcp gene and a wild-type copy of its promoter or promoter with the inverted repeat region deleted. Both strains were then grown in 2 mM nitrite. Data are representative of 3 biological replicates.

To further explore the HcpR binding site, a plasmid-complemented strain was utilized. The Pg108-hcp plasmid is capable of rescuing the nitrite-sensitive phenotype of the V3239 strain by expressing a complete copy of hcp (Fig. 3B). To confirm the HcpR binding site, the inverted repeat sequence in the hcp promoter was removed via PCR mutagenesis. The plasmid containing the wild-type promoter and hcp gene is capable of complementing the loss of the V3239 knockout strain. However, the plasmid without the inverted repeat is not capable of rescuing the knockout strain. This indicates that the inverted repeat is necessary for proper regulation of the hcp gene and is most likely the binding site of HcpR.

P. gingivalis hcp mutant has decreased survival with host cells.Our previous studies have shown that the V2807 HcpR-deficient strain has a reduced ability to survive with host cells (12). To determine if this survival is mediated by Hcp as well as to define the role of Hcp in P. gingivalis interaction with host cells, we investigated the ability of V3239 to survive with human oral keratinocyte (HOK) cells (28). Because the activity of NOS enzymes requires oxygen, all host cell interaction experiments were performed in 6% O2 for 1 h (29). Although it is an anaerobe, P. gingivalis is capable of robust growth under microaerophilic conditions (30, 31). As shown in Fig. 4, the wild-type strain was recovered efficiently after interaction with host cells. We recovered significantly less of the V3282 hcp deletion strain after exposure to HOK cells. Complementation of V3282 resulted in a partial rescue of the wild-type phenotype. Furthermore, wild-type colonies began to appear on plates approximately 4 to 5 days after the experiment; however, the V3282 strain took 7 to 8 days for colonies to appear. These results indicate that Hcp plays an important role in the survival and viability of P. gingivalis exposed to host cells.

FIG 4
FIG 4

Hcp is required for survival of P. gingivalis with host cells. A host cell interaction assay was performed using wild-type (W83), hcp-deficient (V3282), and complemented (V3284) P. gingivalis strains. HOKs were infected with bacteria at an MOI of 200:1 for 1 h. After infection, the host cells were lysed and the bacteria plated on TSA blood agar plates. Data from 3 biological replicates performed in duplicate are represented. (A) Total bacteria recovered from the host cell interaction assay. Cells were washed with PBS and lysed to release bacteria. (B) Intracellular bacteria recovered from the host cell interaction assay. Host cells were treated with 300 μg/ml metronidazole and 400 μg/ml gentamicin for 1 h to eliminate extracellular bacteria and washed. Host cells were lysed to release intracellular bacteria. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

To determine if association or entry of the hcp mutant strain into host cells was compromised, we performed confocal microscopy. P. gingivalis strains were labeled using 2 μM 2′,7′-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein and acetoxymethyl ester (BCECF-AM), and HOKs were observed after infection using 4′,6-diamidino-2-phenylindole (DAPI) and differential interference contrast (DIC) imaging (Fig. 5A and B and Fig. S2). There was no difference in the number of HOK cells containing the wild-type or hcp mutant P. gingivalis strains (either bound to the HOKs or internalized) (Fig. 5C). This indicates that Hcp is important for the survival of the bacterium in the presence of host cells but that its absence does not compromise entry of P. gingivalis into or association with host cells. Thus, the higher recovery of the wild-type strain is due to an Hcp-mediated ability to survive and respond to stresses exhibited by the host cells.

FIG 5
FIG 5

Hcp is not required for binding and internalization of P. gingivalis into host cells. P. gingivalis was labeled with BCECF-AM (green), nuclei were stained with DAPI (blue), and HOKs were visualized via DIC using a confocal microscope. (A) HOK infection with P. gingivalis W83. (B) HOK infection with V3282-deficient P. gingivalis. (C) Infection rate was quantified by counting the number of HOK cells associated with P. gingivalis (intracellular or attached) in 5 to 10 random areas (approximately 100 cells). Images are representative of 3 biological replicates.

DISCUSSION

Periodontal disease is one of the most prevalent diseases in the world, and P. gingivalis plays a significant role in its development and progression. Understanding the ability of P. gingivalis to adapt and survive in the oral cavity, where it encounters high levels of reactive nitrogen, has practical applications. Vegetables and fruits that are high in nitrates are recommended for daily consumption, and they may also have added benefits to oral health. When these foods are eaten they drastically increase the levels of nitrite found in the oral cavity, which can promote the growth of health-promoting commensal bacteria found in the oral cavity, such as Neisseria flavescens and related species (7). P. gingivalis, however, may also withstand high levels of nitrite; thus, mechanisms that would interfere with such resistance may have an added benefit of impairing P. gingivalis growth and colonization via nitrite stress.

Our RNA-seq data show that hcp is the most upregulated gene in P. gingivalis exposed to nitrite. Concentrations as low as 200 μM nitrite result in a 140- to 200-fold upregulation of hcp. These results are supported by the qRT-PCR data. Furthermore, nanomolar amounts of NO are capable of efficiently stimulating the expression of hcp. The upregulation of hcp is abrogated in the HcpR-deficient strain V2807. Furthermore, with hcp being upregulated by large margins, over 140-fold, while few other genes had more than 2-fold upregulation and there were no significantly downregulated genes, HcpR appears most likely to act as a transcriptional activator. The low thresholds indicate that HcpR is very sensitive to reactive nitrogen species it encounters in the environment. The response time is also rapid and prolonged. A more than 200-fold increase of hcp transcript can be achieved in under 15 min, and the increased transcription of hcp persists for more than 1 h after initial exposure of 200 μM NO2−. Under these conditions, it appears that HcpR does not regulate any gene apart from hcp, which implies that the role of HcpR is to efficiently and abruptly regulate hcp in response to reactive nitrogen species.

Of note, when we compared the transcriptional profiles of the V2807 strain grown with and without nitrite, two of the most upregulated genes code for both subunits of the cytochrome d ubiquinol oxidase, cydA and cydB. Under microaerophilic conditions, P. gingivalis has been shown to express and utilize the cydAB operon to consume oxygen and mediate oxygen-based metabolism (30). Studies in the anaerobic bacteria Moorella thermoacetica reveal that cytochrome bd ubiquinol oxidase helps to protect against oxidative stress and contributes to oxygen tolerance (32). Reactive nitrogen species such as NO can react with reactive oxygen species such as superoxide. This reaction creates the very reactive peroxynitrite anion (33). It is possible that P. gingivalis is attempting to avoid the production of these products by reducing the levels of any reactive oxygen species in the environment. However, this is seemingly a secondary effect of HcpR inactivation, and the cyd operon may not be regulated directly by HcpR. The lack of an hcpR binding site in the promoter region supports this hypothesis.

The nrfAH nitrite reductase system has been shown to be upregulated in response to low or intermediate levels of nitrite in an anaerobic environment in E. coli and Desulfovibrio spp. (20, 25). However, in our RNA-seq studies we did not see any change in the P. gingivalis homologues of nrfAH (PG1820 and PG1821) in response to nitrite. Furthermore, we did not see regulation of any of the genes suspected to play a role in nitrite reduction (such as PG2213). Although nrfAH mutants had a decrease in growth rate with respect to the wild-type strain, this was not reliant on nitrite. This implies that their transcription is not fully dependent on nitrite, and this locus may play no role in protecting the bacteria from nitrite. Also, its activity may be regulated at the translational or protein level, which was not investigated in this study.

We show that growth of the hcp-deficient strain is greatly reduced relative to that of the wild-type strain in physiological concentrations of nitrite. As a pathogenic bacterium, P. gingivalis is capable of adhering to, invading, and multiplying inside host cells. To survive the host immune response, P. gingivalis must employ protective mechanisms against host-derived stresses such as oxidative and nitrosative species. We show that the hcp-deficient strain has a greatly reduced capacity to survive with host cells. These studies, combined with the decrease of growth in the presence of nitrite, establish Hcp as biologically significant and necessary for survival in the oral cavity. Furthermore, this work substantiates previous studies on the biological relevance and importance of Hcp and the HcpR regulator (11, 13).

Despite the importance of Hcp in nitrogen-based metabolism and in the response to reactive nitrogen species in obligate and facultative anaerobes, the mechanisms of gene regulation are only now being elucidated. The NsrR and FNR regulators have been shown to regulate hcp in E. coli and other related species (34, 35). Furthermore, recent studies have also implicated the redox-sensitive regulator OxyR in the regulation of hcp (36). However, the role that these regulators play in hcp expression is not consistent with those found in many Gram-negative obligate anaerobes (specifically of the Bacteroides and Desulfovibrio phyla and related anaerobic species) (37, 38). In P. gingivalis, OxyR has no observable effect on hcp expression and regulation (11). Thus, this and previous studies imply that HcpR is the primary regulator of hcp expression.

The hcp null strain has a phenotype very similar to that of the hcpR null strain and does not grow in similar concentrations of nitrite (11). We show that HcpR directly regulates the expression of hcp via binding to the promoter region. Deleting the inverted repeat sequence in the hcp promoter results in a phenotype similar to that of the Hcp-deficient strain, indicating that HcpR regulates hcp expression via directly binding to the hcp promoter at an inverted repeat region upstream of the transcription start site. This agrees with the observed paradigm of FNR-CRP regulators binding to inverted repeats and acting as transcriptional activators (39–41).

hcp encodes a putative NO reductase, although the gene product’s exact function(s) in P. gingivalis is still under much scrutiny, with other studies claiming that it may function as a hydroxylamine reductase (16). In other obligate and facultative anaerobes, Hcp has been shown to play an important role in NO and hydroxylamine reduction and detoxification; however, in the case of P. gingivalis, as shown in our studies, hydroxylamine does not stimulate hcp expression. Recent studies have implicated Hcp as a regulator of protein S-nitrosylation and signaling in E. coli, where it plays a crucial role in the regulation of nitrogen metabolism and the response to by-products of nitrogen metabolism (19). We show that hcp is necessary for growth in nitrite. It is possible that it functions directly in a detoxification role using its enzymatic activity; however, it may also work to posttranslationally modify and regulate other components of the nitrosative stress response, as similarly shown for the E. coli Hcp. Further studies will be required to fully define Hcp’s role in P. gingivalis and its mechanisms of action.

Many nitrosative stress response genes have been identified in bacteria, and these genes have been shown to be regulated by a wide range of regulators, such as NsrR, OxyR, FNR, and HcpR (34, 42, 43). Many bacteria employ a multifactorial approach to respond to nitrosative stress, utilizing a combination of reductase, redox-sensitive iron proteins, and globins to reduce toxic levels of reactive nitrogen species. A transcriptomics study in the anaerobic bacterium Desulfovibrio vulgaris reveals that the bacterium employs multiple pathways in response to nitrite stress, including hcp, the cytochrome c reductase nrfAH operon, NirT cytochrome c protein, and many putative redox-sensitive iron sulfur proteins (44, 45). The relationship between E. coli and nitrate-nitrite reduction has also been well documented. E. coli encodes three distinct nitrate reductase systems and two separate nitrite reductase systems, as well as multiple nitrate-nitrite transporters (4, 46–48). Furthermore, E. coli also employs several enzymes, such as hmp (flavohemoglobin), hcr, and hcp, to coordinate the response to the reactive species produced by nitrate and nitrite reduction, making it well equipped to deal with any nitrosative stress (34, 43). Compared to that of E. coli, P. gingivalis’ transcriptional response to nitrite, or lack thereof, is somewhat unexpected. P. gingivalis appears to be largely dependent on Hcp (and HcpR) to respond to nitrite and NO toxicity under the conditions observed. While Hcp performs this role ably, there appears to be no fallback should Hcp function be compromised.

In summary, this work establishes the biological relevance of hcp and verifies the direct regulation of hcp by the HcpR transcriptional regulator. We show that hcp is necessary for growth in physiological concentrations of nitrite and for survival with host cells. In addition, the transcriptional response of P. gingivalis to nitrite was reported and analyzed.

MATERIALS AND METHODS

Bacterial strains and growth conditions.All strains of P. gingivalis are derived from the W83 type strain and are listed in Table S1 in the supplemental material. All bacteria were grown anaerobically in an atmosphere consisting of 85% N2, 10% CO2, and 5% H2 at 37°C in an anaerobic chamber (Coy Manufacturing). All P. gingivalis strains were maintained on tryptic soy agar (TSA; with sheep’s blood) plates (Fisher). Overnight inoculations of liquid cultures were grown in brain heart infusion (BHI) broth supplemented with hemin (5.0 μg/ml) and vitamin K3 (1 μg/ml).

Growth studies.Overnight cultures of P. gingivalis grown in BHI broth were started from TSA blood agar plates and allowed to grow to stationary phase (24 to 48 h). Consistent with our previous work, mycoplasma broth was utilized in this study (12). The overnight culture served to inoculate samples in mycoplasma broth supplemented with vitamin K3 (1 μg/ml). All samples were started at an optical density at 660 nm (OD660) of 0.10, and growth was monitored at time points of 3, 6, and 24 h or 2, 4, 6, and 24 h. For growth studies and exposure studies in nitrite, cultures were prepared in mycoplasma broth with the appropriate concentration of sodium nitrite or S-nitrosoglutathione (GSNO).

Conditioned medium growth studies were performed in mycoplasma broth. Before growth studies, wild-type P. gingivalis was grown to mid-log phase (OD660 of 0.5 to 0.7) in 2 mM nitrite. The culture was then spun down and sterilized through a 0.22-μm filter. The conditioned medium was diluted 1:2 with fresh mycoplasma and then utilized in growth studies.

Generation of hcp (PG0893), nrfA (PG1821), and nrfH (PG1821) mutant strains.The nrfH (PG1820), nrfA (PG1821), and hcp (PG0893) deletion constructs were created by replacing the coding region of the genes with an ermF cassette. Thus, the ermF cassette and 300 bp upstream and downstream of the protein coding region, including 15-bp overlaps, were PCR amplified. The 3 fragments were assembled into the PCR-2.1 vector using the NEB HiFi assembly master mix (New England Biolabs). The assembled constructs were then transformed and screened in E. coli DH5α, and positive clones were subsequently sequenced.

Constructs for the deletion of hcp, PG1820, and PG1821 were amplified by PCR using gene-specific primers and concentrated to approximately 1 μg/μl. Five μl of the concentrated PCR product was then added to 50 μl of electrocompetent P. gingivalis cells and placed into an electroporation cuvette. The samples were electroporated using a Gene Pulser II electroporation system (Bio-Rad). Immediately after electroporation, 500 μl of warm BHI broth was added to the cuvette and immediately placed in the anaerobic chamber, where 550 μl of sample was added to 2 ml of anaerobic BHI broth. After growing overnight, the samples were then plated on TSA blood agar plates with 0.5 μg/ml of clindamycin for selection. Colonies appeared after approximately 1 week and were replated. To screen the colonies, PCR was performed on the sample using gene-specific primers, and the PCR products were sequenced to confirm the desired genetic makeup of the strains. Strains positive for the replacement mutations were stored and are listed in Table S1.

Generation of complemented strains.Two types of hcp complementation were done. First, genomic complementation with an intact copy of the hcp gene was electroporated into the hcp-deficient strain as described previously (12). Revertants were then selected by plating on blood agar plates containing 2 mM nitrite. Colonies were selected and screened for the reversion and passaged in BHI plus nitrite. The revertants were verified by the absence of growth in 0.5 μg/ml clindamycin, and their genetic makeup was confirmed by PCR followed by DNA sequencing.

A plasmid complementation also was done. The Pg108 vector is a shuttle vector derived from the pYHBA1 plasmid and confers tetracycline resistance in P. gingivalis and erythromycin resistance in E. coli (49). The hcp gene and its native promoter were amplified using primers Pg108-hcp-F and Pg108-hcp-R (Table S2). The amplified DNA was cloned into the Pg108 vector. The assembled vector was transformed and cloned in E. coli DH5α (New England Biolabs). The sequenced vector was electroporated into the hcp-deficient strain (V3239) as described above. All plasmids used in this study are listed in Table S3.

ChIP assay with HcpR.An HcpR construct that places a 3× FLAG tag on the N terminus of HcpR downstream of the ermF promoter was synthesized and cloned into the Pg108 vector at the SphI and BamHI sites to create the Pg108-hcpR-FLAG plasmid (Table S3). To generate a truncated version of HcpR, this plasmid was mutated using the L156*-Forward and L156*-Reverse primers using the QuikChange site-directed mutagenesis kit (Agilent Technologies) to create the plasmid Pg108-L156*-hcpR-FLAG (Table S3). This plasmid expresses a truncated FLAG-tagged HcpR lacking the DNA binding domain and was used as a negative control. Both of these plasmids were electroporated into the hcpR-deficient mutant strain (V2807) to create the strains V3243 (Pg108-hcpR-FLAG) and V3237 (Pg108-hcpR-FLAG L156*). Strains V3243 and V3237 were grown on blood plates with appropriate antibiotics and used to inoculate an overnight culture of BHI medium. The overnight culture was used to inoculate a culture of 5 ml mycoplasma medium with 0.25 μg/ml tetracycline at an OD660 of 0.15. At mid-log phase the cultures were exposed to 2 mM nitrite for 15 min. After nitrite treatment, formaldehyde was added to a final concentration of 1% for cross-linking and was incubated at room temperature for 20 min. After incubation, cross-linking was quenched via the addition of glycine to a final concentration of 0.5 M. Cells were harvested via centrifugation and washed with ice-cold phosphate-buffered saline (PBS). Washed cells were suspended in 1 ml of lysis buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1%-Triton X-100, protease inhibitor cocktail). Lysozyme was added to a final concentration of 1 mg/ml along with CelLytic B (Sigma) to lyse cells. Cells were lysed at room temperature for 15 min. After incubation, the lysis mixture was sheared on ice in a bath sonicator. After sonication, insoluble material was removed via centrifugation at 13,000 × g for 10 min. The cleared lysates were added to anti-FLAG M2 magnetic beads (Sigma-Aldrich). The samples were incubated for 2 h at room temperature with gentle shaking. Resin was washed 6 times at room temperature with 20× packed gel volumes of Tris-buffered saline (TBS) on a magnetic separator. Samples were eluted from the resin using 5 packed gel volumes of 3× FLAG peptide elution buffer (TBS containing 20 μg/ml of FLAG peptide). Samples were incubated with elution buffer for 30 min. After incubation, the supernatant was removed and stored at −20°C. Cross-linking was reversed via treatment with 20 μg protease K at 65°C for 15 min and then boiled for 15 min. PCR utilizing primers specific for the hcp promoter was performed to probe for the presence of the promoter region in the elutions. Primers are listed in Table S2.

RNA-seq library generation.Samples of P. gingivalis W83 and V2807 (hcpR mutant) were grown overnight in BHI medium. The overnight cultures served to inoculate mycoplasma medium starting at an OD660 of approximately 0.10. Cultures were grown to mid-log phase and then exposed to 200 μM nitrite for 1 h before being centrifuged and stored at −20°C. RNA isolation was carried out using the RNeasy minikit (Qiagen) by following the manufacturer’s protocol. Residual DNA was removed using the DNA-free DNase kit (Ambion) by following the manufacturer’s protocol. An agarose gel was run on each sample to confirm sample fidelity. For RNA-seq library generation, the Ovation complete prokaryotic RNA-seq DR multiplex kit (Nugen) was used. The library was generated by following the manufacturer’s protocol, and fragment sizes from sonication during library generation were confirmed using a bioanalyzer (Agilent Technologies). Libraries were sequenced by the VCU nucleic acid sequencing core via MiSeq Illumina sequencing. Sequence reads were aligned to the genome of P. gingivalis W83 using CLC Genomics Workbench (CLC Bio). Differential gene expression was determined by comparing the number of reads/gene for W83 and V2807 with and without nitrite. For the comparison between samples, any gene with fewer than 10 reads was not analyzed, and genes that were differentially regulated ∼2.0-fold were selected for further analysis.

qRT-PCR analysis of gene expression.Real-time qRT-PCR was done with a SYBR green-based detection system on an Applied Biosystems 7500 Fast real-time PCR system (Applied Biosystems). Briefly, cells were grown to mid-log phase and exposed to reactive nitrogen species as detailed above. The RNeasy minikit (Qiagen) was used to purify RNA from cells, and residual DNA was removed using the DNA-free DNase kit (Ambion) by following the manufacturer’s protocol. The cDNA was generated using a cDNA synthesis kit (Applied Biosystems) by following the manufacturer’s protocol. Quantitative PCR was performed using 10 ng of cDNA and the ΔΔCT method (where CT is threshold cycle), utilizing the 16S ribosomal subunit as an endogenous control. Primers used in this study are listed in Table S2.

Dot blot analysis.An Hcp construct that places a 3× FLAG tag at the N terminus of Hcp downstream of its native promoter was synthesized and cloned into the Pg108 vector at the PstI and BamHI sites to create the Pg108-hcp-FLAG plasmid (Table S3). This plasmid was electroporated into the P. gingivalis hcp mutant (V3239) to create strain V3290. The Hcp-FLAG tag-complemented strain was exposed to nitrite for 1 h in mycoplasma broth. After exposure, the extracellular fraction was saved and the cells were lysed. Ten μg of total protein was blotted to a nitrocellulose membrane. The membrane was blocked using 3% milk, and FLAG-specific monoclonal antibodies (Sigma) were added to the membrane. The blots were imaged using peroxidase-conjugated goat anti-mouse IgG (Jackson Immuno Research) on film.

Host cell interaction assay.HOKs (ScienCell Research Laboratories) were grown to confluence in flasks and plated in 12-well tissue culture plates in a cell-specific medium. The cells were infected with wild-type P. gingivalis, hcp mutant strain V3282, or the complemented strain V3284 at a multiplicity of infection (MOI) of 200:1. Infection was conducted for 1 h under an atmosphere of 6% oxygen at 37°C. After infection, cells in each well were washed in an anaerobic chamber 3 times using 1 ml PBS. After washing, cells were lysed using 1% saponin to release intracellular bacteria. Lysed cells were serially diluted using anaerobic BHI medium and plated on TSA blood agar plates. Colonies appeared after 5 to 8 days and were counted to calculate the number of CFU/ml for each infection.

Confocal microscopy.HOK cells were seeded and grown overnight on poly-l-lysine-treated 18-mm circular cover-glass slides. Prior to infection, P. gingivalis strains were incubated with BCECF-AM for 30 min in PBS. After incubation, the bacteria were pelleted and resuspended to remove excess BCECF-AM. HOK cells were infected at an MOI of 200:1 for 1 h under an atmosphere of 6% oxygen at 37°C. After infection, cells were washed 3 times using PBS. HOKs were fixed using 4% formaldehyde in PBS for 10 min at room temperature. Cells were permeabilized using 0.1% Triton X-100 for 10 min. Nuclei were stained using DAPI (Vector Laboratories) while mounting coverslips on microscope slides. For determination of the infection rate, the number of HOK cells with internalized P. gingivalis was counted and divided by the total number of cells in 5 to 10 random areas (approximately 100 cells). Confocal images were acquired on a Zeiss LSM 700 laser scanning microscope.

Data analysis.All experiments were performed at least three times. Data in graphs are presented as means, and error bars are presented as the standard deviations (SD) between replicates. The statistical analysis was performed via paired t test for sample means of the wild type and mutant. A P value of <0.05 was considered statistically significant.

Data availability.RNA-seq data were deposited in the Gene Expression Omnibus (GEO) with the reference number GSE117421.

ACKNOWLEDGMENTS

This work was funded by NIH grants 1R01DE023304 (J.P.L.) and 1F31DE025158 (B.R.B.). Microscopy was performed at the VCU microscopy facility, supported in part by funding from the NIH-NCI Cancer Center Support grant P30 CA016059.

B.R.B. and J.P.L. designed this study; Q.G. and B.R.B. performed host cell experiments; B.R.B. and J.A.H. developed mutant strains of P. gingivalis; B.R.B. performed all other experiments. B.R.B. and J.P.L. analyzed all data, and B.R.B. wrote the initial manuscript. All authors contributed to writing and editing the final manuscript.

FOOTNOTES

    • Received 27 July 2018.
    • Returned for modification 23 August 2018.
    • Accepted 11 January 2019.
    • Accepted manuscript posted online 22 January 2019.

  • Supplemental material for this article may be found at https://doi.org/10.1128/IAI.00572-18.

REFERENCES

  1. 1.↵
    1. Hajishengallis G,
    2. Darveau RP,
    3. Curtis MA
    . 2012. The keystone-pathogen hypothesis. Nat Rev Microbiol 10:717–725. doi:10.1038/nrmicro2873.
    OpenUrlCrossRefPubMed
  2. 2.↵
    1. Mysak J,
    2. Podzimek S,
    3. Sommerova P,
    4. Lyuya-Mi Y,
    5. Bartova J,
    6. Janatova T,
    7. Prochazkova J,
    8. Duskova J
    . 2014. Porphyromonas gingivalis: major periodontopathic pathogen overview. J Immunol Res 2014:1. doi:10.1155/2014/476068.
    OpenUrlCrossRef
  3. 3.↵
    1. Weitzberg E,
    2. Lundberg JO
    . 1998. Nonenzymatic nitric oxide production in humans. Nitric Oxide 2:1–7. doi:10.1006/niox.1997.0162.
    OpenUrlCrossRefPubMedWeb of Science
  4. 4.↵
    1. Lundberg JO,
    2. Weitzberg E,
    3. Cole JA,
    4. Benjamin N
    . 2004. Nitrate, bacteria and human health. Nat Rev Microbiol 2:593–602. doi:10.1038/nrmicro929.
    OpenUrlCrossRefPubMedWeb of Science
  5. 5.↵
    1. Lundberg JO,
    2. Weitzberg E,
    3. Gladwin MT
    . 2008. The nitrate-nitrite-nitric oxide pathway in physiology and therapeutics. Nat Rev Drug Discov 7:156–167. doi:10.1038/nrd2466.
    OpenUrlCrossRefPubMedWeb of Science
  6. 6.↵
    1. Hezel MP,
    2. Weitzberg E
    . 2015. The oral microbiome and nitric oxide homoeostasis. Oral Dis 21:7–16. doi:10.1111/odi.12157.
    OpenUrlCrossRefPubMed
  7. 7.↵
    1. Velmurugan S,
    2. Gan JM,
    3. Rathod KS,
    4. Khambata RS,
    5. Ghosh SM,
    6. Hartley A,
    7. Van Eijl S,
    8. Sagi-Kiss V,
    9. Chowdhury TA,
    10. Curtis M,
    11. Kuhnle GGC,
    12. Wade WG,
    13. Ahluwalia A
    . 2016. Dietary nitrate improves vascular function in patients with hypercholesterolemia: a randomized, double-blind, placebo-controlled study. Am J Clin Nutr 103:25–38. doi:10.3945/ajcn.115.116244.
    OpenUrlAbstract/FREE Full Text
  8. 8.↵
    1. Alderton WK,
    2. Cooper CE,
    3. Knowles RG
    . 2001. Nitric oxide synthases: structure, function and inhibition. Biochem J 357:593–615. doi:10.1042/bj3570593.
    OpenUrlAbstract/FREE Full Text
  9. 9.↵
    1. Gyurko R,
    2. Shoji H,
    3. Battaglino RA,
    4. Boustany G,
    5. Gibson FC,
    6. Genco CA,
    7. Stashenko P,
    8. Van Dyke TE
    . 2005. Inducible nitric oxide synthase mediates bone development and P. gingivalis-induced alveolar bone loss. Bone 36:472–479. doi:10.1016/j.bone.2004.12.002.
    OpenUrlCrossRefPubMed
  10. 10.↵
    1. Herrera BS,
    2. Martins-Porto R,
    3. Maia-Dantas A,
    4. Campi P,
    5. Spolidorio LC,
    6. Costa SK,
    7. Van Dyke TE,
    8. Gyurko R,
    9. Muscara MN
    . 2011. iNOS-derived nitric oxide stimulates osteoclast activity and alveolar bone loss in ligature-induced periodontitis in rats. J Periodontol 82:1608–1615. doi:10.1902/jop.2011.100768.
    OpenUrlCrossRefPubMed
  11. 11.↵
    1. Fang FC
    . 2004. Antimicrobial reactive oxygen and nitrogen species: concepts and controversies. Nat Rev Microbiol 2:820–832. doi:10.1038/nrmicro1004.
    OpenUrlCrossRefPubMedWeb of Science
  12. 12.↵
    1. Lewis JP,
    2. Yanamandra SS,
    3. Anaya-Bergman C
    . 2012. HcpR of Porphyromonas gingivalis is required for growth under nitrosative stress and survival within host cells. Infect Immun 80:3319–3331. doi:10.1128/IAI.00561-12.
    OpenUrlAbstract/FREE Full Text
  13. 13.↵
    1. Miller DP,
    2. Hutcherson JA,
    3. Wang Y,
    4. Nowakowska ZM,
    5. Potempa J,
    6. Yoder-Himes DR,
    7. Scott DA,
    8. Whiteley M,
    9. Lamont RJ
    . 2017. Genes contributing to Porphyromonas gingivalis fitness in abscess and epithelial cell colonization environments. Front Cell Infect Microbiol 7:378. doi:10.3389/fcimb.2017.00378.
    OpenUrlCrossRef
  14. 14.↵
    1. Boutrin MC,
    2. Wang C,
    3. Aruni W,
    4. Li X,
    5. Fletcher HM
    . 2012. Nitric oxide stress resistance in porphyromonas gingivalis is mediated by a putative hydroxylamine reductase. J Bacteriol 194:1582–1592. doi:10.1128/JB.06457-11.
    OpenUrlAbstract/FREE Full Text
  15. 15.↵
    1. Rodionov DA,
    2. Dubchak I,
    3. Arkin A,
    4. Alm E,
    5. Gelfand MS
    . 2004. Reconstruction of regulatory and metabolic pathways in metal-reducing delta-proteobacteria. Genome Biol 5:R90. doi:10.1186/gb-2004-5-11-r90.
    OpenUrlCrossRefPubMed
  16. 16.↵
    1. Wang J,
    2. Vine CE,
    3. Balasiny BK,
    4. Rizk J,
    5. Bradley CL,
    6. Tinajero-Trejo M,
    7. Poole RK,
    8. Bergaust LL,
    9. Bakken LR,
    10. Cole JA
    . 2016. The roles of the hybrid cluster protein, Hcp and its reductase, Hcr, in high affinity nitric oxide reduction that protects anaerobic cultures of Escherichia coli against nitrosative stress. Mol Microbiol 100:877–892. doi:10.1111/mmi.13356.
    OpenUrlCrossRef
  17. 17.↵
    1. Cadby IT,
    2. Faulkner M,
    3. Cheneby J,
    4. Long J,
    5. Van Helden J,
    6. Dolla A,
    7. Cole JA
    . 2017. Coordinated response of the Desulfovibrio desulfuricans 27774 transcriptome to nitrate, nitrite and nitric oxide. Sci Rep 7:16228. doi:10.1038/s41598-017-16403-4.
    OpenUrlCrossRef
  18. 18.↵
    1. Wolfe MT,
    2. Heo J,
    3. Garavelli JS,
    4. Ludden PW
    . 2002. Hydroxylamine reductase activity of the hybrid cluster protein from Escherichia coli. J Bacteriol 184:5898–5902. doi:10.1128/JB.184.21.5898-5902.2002.
    OpenUrlAbstract/FREE Full Text
  19. 19.↵
    1. Seth D,
    2. Hess DT,
    3. Hausladen A,
    4. Wang L,
    5. Wang Y,
    6. Stamler JS
    . 2018. A multiplex enzymatic machinery for cellular protein S-nitrosylation. Mol Cell 69:451–464.e6. doi:10.1016/j.molcel.2017.12.025.
    OpenUrlCrossRef
  20. 20.↵
    1. Wang H,
    2. Gunsalus RP
    . 2000. The nrfA and nirB nitrite reductase operons in Escherichia coli are expressed differently in response to nitrate than to nitrite. J Bacteriol 182:5813–5822. doi:10.1128/JB.182.20.5813-5822.2000.
    OpenUrlAbstract/FREE Full Text
  21. 21.↵
    1. Cole J
    . 1996. Nitrate reduction to ammonia by enteric bacteria: redundancy, or a strategy for survival during oxygen starvation? FEMS Microbiol Lett 136:1–11. doi:10.1111/j.1574-6968.1996.tb08017.x.
    OpenUrlCrossRefPubMedWeb of Science
  22. 22.↵
    1. Meuric V,
    2. Rouillon A,
    3. Chandad F,
    4. Bonnaure-Mallet M
    . 2010. Putative respiratory chain of Porphyromonas gingivalis. Future Microbiol 5:717–734. doi:10.2217/fmb.10.32.
    OpenUrlCrossRefPubMed
  23. 23.↵
    1. Kern M,
    2. Volz J,
    3. Simon J
    . 2011. The oxidative and nitrosative stress defence network of Wolinella succinogenes: cytochrome c nitrite reductase mediates the stress response to nitrite, nitric oxide, hydroxylamine and hydrogen peroxide. Environ Microbiol 13:2478–2494. doi:10.1111/j.1462-2920.2011.02520.x.
    OpenUrlCrossRefPubMed
  24. 24.↵
    1. Pittman MS,
    2. Elvers KT,
    3. Lee L,
    4. Jones MA,
    5. Poole RK,
    6. Park SF,
    7. Kelly DJ
    . 2007. Growth of Campylobacter jejuni on nitrate and nitrite: electron transport to NapA and NrfA via NrfH and distinct roles for NrfA and the globin Cgb in protection against nitrosative stress. Mol Microbiol 63:575–590. doi:10.1111/j.1365-2958.2006.05532.x.
    OpenUrlCrossRefPubMed
  25. 25.↵
    1. Rajeev L,
    2. Chen A,
    3. Kazakov AE,
    4. Luning EG,
    5. Zane GM,
    6. Novichkov PS,
    7. Wall JD,
    8. Mukhopadhyay A
    . 2015. Regulation of nitrite stress response in Desulfovibrio vulgaris Hildenborough, a model sulfate-reducing bacterium. J Bacteriol 197:3400–3408. doi:10.1128/JB.00319-15.
    OpenUrlAbstract/FREE Full Text
  26. 26.↵
    1. Broniowska KA,
    2. Diers AR,
    3. Hogg N
    . 2013. S-Nitrosoglutathione. Biochim Biophys Acta 1830:3173–3181. doi:10.1016/j.bbagen.2013.02.004.
    OpenUrlCrossRefPubMed
  27. 27.↵
    1. Noble DR,
    2. Swift HR,
    3. Williams DLH
    . 1999. Nitric oxide release from S-nitrosoglutathione (GSNO). Chem Commun (Camb):2317–2318. doi:10.1039/a907891f.
    OpenUrlCrossRef
  28. 28.↵
    1. Lamont RJ,
    2. Chan A,
    3. Belton CM,
    4. Izutsu KT,
    5. Vasel D,
    6. Weinberg A
    . 1995. Porphyromonas gingivalis invasion of gingival epithelial cells. Infect Immun 63:3878–3885.
    OpenUrlAbstract/FREE Full Text
  29. 29.↵
    1. Griffi OW,
    2. Stuehr DJ
    . 1995. Nitric oxide synthases: properties and catalytic mechanism. Annu Rev Physiol 57:707–736. doi:10.1146/annurev.ph.57.030195.003423.
    OpenUrlCrossRefPubMedWeb of Science
  30. 30.↵
    1. Lewis JP,
    2. Iyer D,
    3. Anaya-Bergman C
    . 2009. Adaptation of Porphyromonas gingivalis to microaerophilic conditions involves increased consumption of formate and reduced utilization of lactate. Microbiology 155:3758–3774. doi:10.1099/mic.0.027953-0.
    OpenUrlCrossRefPubMedWeb of Science
  31. 31.↵
    1. He J,
    2. Miyazaki H,
    3. Anaya C,
    4. Yu F,
    5. Yeudall WA,
    6. Lewis JP
    . 2006. Role of Porphyromonas gingivalis FeoB2 in metal uptake and oxidative stress protection. Infect Immun 74:4214–4223. doi:10.1128/IAI.00014-06.
    OpenUrlAbstract/FREE Full Text
  32. 32.↵
    1. Das A,
    2. Silaghi-Dumitrescu R,
    3. Ljungdahl LG,
    4. Kurtz DM
    . 2005. Cytochrome bd oxidase, oxidative stress, and dioxygen tolerance of the strictly anaerobic bacterium Moorella thermoacetica. J Bacteriol 187:2020–2029. doi:10.1128/JB.187.6.2020-2029.2005.
    OpenUrlAbstract/FREE Full Text
  33. 33.↵
    1. Hill BG,
    2. Dranka BP,
    3. Bailey SM,
    4. Lancaster JR,
    5. Darley-Usmar VM
    . 2010 What part of NO don’t you understand? Some answers to the cardinal questions in nitric oxide biology. J Biol Chem 285:19699–19704. doi:10.1074/jbc.R110.101618.
    OpenUrlAbstract/FREE Full Text
  34. 34.↵
    1. Filenko N,
    2. Spiro S,
    3. Browning DF,
    4. Squire D,
    5. Overton TW,
    6. Cole J,
    7. Constantinidou C
    . 2007. The NsrR regulon of Escherichia coli K-12 includes genes encoding the hybrid cluster protein and the periplasmic, respiratory nitrite reductase. J Bacteriol 189:4410–4417. doi:10.1128/JB.00080-07.
    OpenUrlAbstract/FREE Full Text
  35. 35.↵
    1. Filenko NA,
    2. Browning DF,
    3. Cole JA
    . 2005. Transcriptional regulation of a hybrid cluster (prismane) protein. Biochem Soc Trans 33:195–197. doi:10.1042/BST0330195.
    OpenUrlAbstract/FREE Full Text
  36. 36.↵
    1. Seth D,
    2. Hausladen A,
    3. Wang YJ,
    4. Stamler JS
    . 2012. Endogenous protein S-nitrosylation in E. coli: regulation by OxyR. Science 336:470–473. doi:10.1126/science.1215643.
    OpenUrlAbstract/FREE Full Text
  37. 37.↵
    1. Rodionov DA,
    2. Dubchak IL,
    3. Arkin AP,
    4. Alm EJ,
    5. Gelfand MS
    . 2005. Dissimilatory metabolism of nitrogen oxides in bacteria: comparative reconstruction of transcriptional networks. PLoS Comp Biol 1:e55. doi:10.1371/journal.pcbi.0010055.eor.
    OpenUrlCrossRef
  38. 38.↵
    1. Cadby IT,
    2. Busby SJ,
    3. Cole JA
    . 2011. An HcpR homologue from Desulfovibrio desulfuricans and its possible role in nitrate reduction and nitrosative stress. Biochem Soc Trans 39:224–229. doi:10.1042/BST0390224.
    OpenUrlAbstract/FREE Full Text
  39. 39.↵
    1. Spiro S
    . 1994. The FNR family of transcriptional regulators. Antonie Van Leeuwenhoek 66:23–36. doi:10.1007/BF00871630.
    OpenUrlCrossRefPubMedWeb of Science
  40. 40.↵
    1. Korner H,
    2. Sofia HJ,
    3. Zumft WG
    . 2003. Phylogeny of the bacterial superfamily of Crp-Fnr transcription regulators: exploiting the metabolic spectrum by controlling alternative gene programs. FEMS Microbiol Rev 27:559–592. doi:10.1016/S0168-6445(03)00066-4.
    OpenUrlCrossRefPubMedWeb of Science
  41. 41.↵
    1. Berg OG,
    2. von Hippel PH
    . 1988. Selection of DNA binding sites by regulatory proteins. II. The binding specificity of cyclic AMP receptor protein to recognition sites. J Mol Biol 200:709–723. doi:10.1016/0022-2836(88)90482-2.
    OpenUrlCrossRefPubMedWeb of Science
  42. 42.↵
    1. Zheng M,
    2. Aslund F,
    3. Storz G
    . 1998. Activation of the OxyR transcription factor by reversible disulfide bond formation. Science 279:1718–1722. doi:10.1126/science.279.5357.1718.
    OpenUrlAbstract/FREE Full Text
  43. 43.↵
    1. Tucker NP,
    2. Le Brun NE,
    3. Dixon R,
    4. Hutchings MI
    . 2010. There’s NO stopping NsrR, a global regulator of the bacterial NO stress response. Trends Microbiol 18:149–156. doi:10.1016/j.tim.2009.12.009.
    OpenUrlCrossRefPubMedWeb of Science
  44. 44.↵
    1. He Q,
    2. Huang KH,
    3. He Z,
    4. Alm EJ,
    5. Fields MW,
    6. Hazen TC,
    7. Arkin AP,
    8. Wall JD,
    9. Zhou J
    . 2006. Energetic consequences of nitrite stress in Desulfovibrio vulgaris Hildenborough, inferred from global transcriptional analysis. Appl Environ Microbiol 72:4370–4381. doi:10.1128/AEM.02609-05.
    OpenUrlAbstract/FREE Full Text
  45. 45.↵
    1. Yurkiw MA,
    2. Voordouw J,
    3. Voordouw G
    . 2012. Contribution of rubredoxin:oxygen oxidoreductases and hybrid cluster proteins of Desulfovibrio vulgaris Hildenborough to survival under oxygen and nitrite stress. Environ Microbiol 14:2711–2725. doi:10.1111/j.1462-2920.2012.02859.x.
    OpenUrlCrossRefPubMed
  46. 46.↵
    1. Sparacino-Watkins C,
    2. Stolz JF,
    3. Basu P
    . 2014. Nitrate and periplasmic nitrate reductases. Chem Soc Rev 43:676–706. doi:10.1039/c3cs60249d.
    OpenUrlCrossRefPubMed
  47. 47.↵
    1. Simon J
    . 2002. Enzymology and bioenergetics of respiratory nitrite ammonification. FEMS Microbiol Rev 26:285–309. doi:10.1111/j.1574-6976.2002.tb00616.x.
    OpenUrlCrossRefPubMedWeb of Science
  48. 48.↵
    1. Rabin RS,
    2. Stewart V
    . 1993. Dual response regulators (NarL and NarP) interact with dual sensors (NarX and NarQ) to control nitrate- and nitrite-regulated gene expression in Escherichia coli K-12. J Bacteriol 175:3259–3268. doi:10.1128/jb.175.11.3259-3268.1993.
    OpenUrlAbstract/FREE Full Text
  49. 49.↵
    1. Yoshimoto H,
    2. Takahasi Y,
    3. Kato D,
    4. Umemoto T
    . 1997. Construction of a plasmid vector for transformation of Porphyromonas gingivalis. FEMS Microbiol Lett 152:175–181. doi:10.1111/j.1574-6968.1997.tb10425.x.
    OpenUrlCrossRefPubMedWeb of Science
Back to top
Download PDF
Citation Tools
Print

Alerts
Email

KEYWORDS

Hcp
HcpR
Porphyromonas gingivalis
anaerobe
host-pathogen
nitrite
nitrosative stress
oral microbiology
periodontitis
virulence

Related Articles

Cited By...