CpsY Influences Streptococcus iniae Cell Wall Adaptations Important for Neutrophil Intracellular Survival

Abstract

The ability of a pathogen to evade neutrophil phagocytic killing mechanisms is critically important for dissemination and establishment of a systemic infection. Understanding how pathogens overcome these innate defenses is essential for the development of optimal therapeutic strategies for invasive infections. CpsY is a conserved transcriptional regulator previously identified as an important virulence determinant for systemic infection of Streptococcus iniae. While orthologs of CpsY have been associated with the regulation of methionine metabolism and uptake pathways, CpsY additionally functions in protection from neutrophil-mediated killing. S. iniae does not alter neutrophil phagosomal maturation but instead is able to adapt to the extreme bactericidal environment of a mature neutrophil phagosome, a property dependent upon CpsY. This CpsY-dependent adaptation appears to involve stabilization of the cell wall through peptidoglycan O-acetylation and repression of cellular autolysins. Furthermore, S. iniae continues to be a powerful model for investigation of bacterial adaptations during systemic streptococcal infection.

INTRODUCTION

The degree to which a systemic pathogen can evade or manipulate the host innate immune response is a key factor in determining the severity of a bacterial infection (79). Polymorphonuclear leukocytes (neutrophils) are a primary bactericidal force against invading bacteria and represent a formidable obstacle for pathogens to overcome (8). This is particularly reflected in the susceptibility of patients with inherited neutrophil disorders to severe bacterial infections (53). Neutrophils are efficient at eliminating invading pathogens through both intracellular (71) and extracellular (59) killing mechanisms. Engulfed pathogens suffer a barrage of bactericidal attacks from exposure to reactive oxygen and nitrogen species (47, 71), antimicrobial peptides (26, 67), proteases (13, 69, 72), peptidoglycan-degrading enzymes (57), and metal deprivation (35, 37, 58).

Invasive pathogens have evolved several strategies for evasion of neutrophil killing mechanisms. Pathogens attempt to avoid phagocytic uptake through prevention of opsonin deposition (76) and production of a polysaccharide capsule (50). Engulfed pathogens can disrupt phagosomal maturation, escape into the cytoplasm, or adapt for survival within the mature phagosome (79). Adaptive processes for phagosomal survival occur in part through chemical modification of outer surface structures, including lysinylation of lipid membranes (23, 44, 70), d-alanylation of wall teichoic acids (15, 24), and O-acetylation of N-acetylmuramyl peptidoglycan residues (3, 4, 19).

Streptococcus iniae is an invasive pathogen of aquatic species that causes severe bacteremia and meningoencephalitis (1, 20, 21). The rapid dissemination of S. iniae to systemic sites reflects its ability to survive phagocytic killing by neutrophils (2, 11, 25, 51). CpsY is a conserved transcriptional regulator previously identified as an important virulence determinant for systemic infection of Streptococcus iniae (51), and mutants of cpsY are specifically susceptible to neutrophil-mediated killing in whole blood (2). CpsY is orthologous to the MetR transcriptional regulator of methionine metabolism and uptake pathways in Streptococcus mutans and Streptococcus agalactiae (10, 73). We previously demonstrated that the susceptibility of an S. iniae cpsY deletion mutant (ΔcpsY) to clearance by neutrophil phagocytosis is independent of its role in methionine supply pathways (2). Thus, the CpsY regulon appears to function for S. iniae intracellular survival in neutrophils through an undetermined mechanism.

This work elucidates a possible role for CpsY during S. iniae intracellular survival in neutrophils. S. iniae does not appear to disrupt neutrophil phagosomal maturation; instead, the CpsY regulon protects S. iniae from the bactericidal components contained within a mature phagosome. This protection appears to be mediated in part by modifications to the peptidoglycan cell wall and repression of streptococcal autolysins.

MATERIALS AND METHODS

Bacterial strains, media, and culture conditions.

The parent Streptococcus iniae strain 9117 and a cpsY in-frame deletion mutant (ΔcpsY) used in this study have been described previously (2, 51). All plasmid constructs were propagated using Escherichia coli TOP10 cells (Invitrogen) cultured in Luria-Bertani (LB) medium. Streptococcal strains were cultured in Todd-Hewitt broth (BBL) supplemented with 0.2% yeast extract (BBL) (THY) (52). Antibiotics were added to the medium when required at the following concentrations: kanamycin at 25 μg ml⁻¹ or chloramphenicol at 20 μg ml⁻¹ for E. coli; kanamycin at 500 μg ml⁻¹, chloramphenicol at 3 μg ml⁻¹, or streptomycin at 1 mg ml⁻¹ for S. iniae. Bacteria were routinely cultured overnight, diluted 1:50 in fresh medium, and grown to mid-exponential phase (optical density at 600 nm [OD₆₀₀], 0.225) at 37°C without shaking, unless otherwise stated. Bacterial growth curves were determined in 96-well plates as previously described (2).

Construction of S. iniae overexpression mutants.

The primers 5′murA.PstI (AAAACTGCAGGAGAAGAAGAATAAAGCCTATAGTG) and 3′ murA.ApaI (TTGGGCCCCTAAAGGCCTTTATTTGTCTTTG) were used to amplify an 860-bp fragment containing the MurNAc-l-alanine amidase (murA; Sin.1445). The fragment was inserted into the PstI/ApaI sites of the vector pLZ12-km (29), generating pLZ12-km-murA. The promoter of the hyaluronic acid synthesis operon in Streptococcus pyogenes (18) was PCR amplified using primers 5′ hasApro.BglII (GGAAGATCTCAGATGAAGTTGTACTCCCTG) and 3′hasApro.BamHI (CGCGGATCCGGCACAATTACACCTCTTC). This fragment was inserted into the BglI/BamHI sites in pLZ12-km-murA to generate the final construct, pLZ12-hasApro.murA.

The vector pLZ12-km-hasApro.murA was cut with PstI and ApaI to remove murA, generating pLZ12-km-hasApro. The primers 5′oatA.PstI (TGCACTGCAGCTTTAAACAGGGGATTTATCAAAAATG) and 3′oatA.ApaI (TGGGGGCCCGACCTTATTTGGCCTTATTTTTG) were used to PCR amplify a 1,815-bp fragment containing the peptidoglycan O-acetyltransferase (oatA; Sin.29). This fragment was inserted into the PstI/ApaI sites of pLZ12-km-hasApro to generate the final construct pLZ12-km-hasApro.oatA. pLZ12-km-hasApro.murA was transformed into the wild-type (WT) S. iniae strain, since it is overexpressed in the ΔcpsY mutant, in an attempt to mimic the ΔcpsY mutant phenotype. pLZ12-km-hasApro.oatA was transformed into the ΔcpsY mutant in an effort to complement the mutant for the WT phenotype. S. iniae transformations were performed as previously described (55).

Neutrophil infection assays.

Primary neutrophils were isolated from human whole blood, and infections were performed in either whole blood or neutrophil preparations as previously described (2). For competition assays, neutrophils were infected as before with minor variations. Mid-exponential-phase cultures were adjusted to 1 × 10⁵ CFU ml⁻¹ in phosphate-buffered saline (PBS), from which equal volumes were pooled and inoculated 1:50 (20-μl volume) into 1-ml neutrophil preparations. Samples were incubated at 37°C for 3 h with gentle rotation, followed by serial dilution in water to lyse infected neutrophils and replica plating on THY agar with or without 1,000 μg ml⁻¹ streptomycin for bacterial enumeration. The competitive index (CI) was calculated by dividing the output ratio (mutant/wild type) by the input ratio (mutant/wild type).

Confocal laser scanning fluorescence microscopy.

S. iniae neutrophil infections were performed as described above for 1 h. One hundred microliters of the sample was loaded into a cytology funnel (Thermo), centrifuged onto glass slides for 3 min at 700 rpm in a Cytospin 1 centrifuge (Shandon Elliott), and fixed in 4% paraformaldehyde (Sigma) for 15 min at room temperature. Fixed cells were washed 3 times in PBS and treated for 30 min with 100 μl of permeabilization buffer (0.2% Tween 20 in PBS plus 10% mutanolysin). Cells were washed in PBS and covered for 30 min in blocking buffer (2% cold water fish gelatin [Sigma] in PBS). One hundred microliters of a polyclonal anti-S. iniae antibody was added to the cells for 2 h, followed by washing 3 times in PBS. One hundred microliters of a secondary Alexa Fluor 488-conjugated goat anti-rabbit antibody (Molecular Probes) was added to the cells for 30 min, and again the mixture was washed 3 times with PBS. This procedure was repeated a second time using a monoclonal antibody specific for human polymorphonuclear leukocyte (PMN) lysosome-associated membrane protein 1 (LAMP-1) (CD107A; Santa Cruz Biotechnology) and a secondary Alexa Fluor tetramethyl rhodamine isocyanate-conjugated goat anti-mouse antibody (Molecular Probes), or using a primary goat antibody for staining of human PMN azurophil (primary) granule marker CD63 (Santa Cruz Biotechnology) and a secondary rhodamine-conjugated chicken anti-goat antibody (Santa Cruz Biotechnology). Prepared cells were mounted in ProLong Gold antifade reagent with 4′,6-diamidino-2-phenylindole (Molecular Probes). Colocalization was determined by fluorescence microscopy using a Leica TCS SP5 laser scanning confocal microscope.

Antimicrobial sensitivity assays.

Working concentrations of the following antimicrobials were prepared in THY broth: colistin (500 μg ml⁻¹; Sigma), gramicidin (1.5 μg ml⁻¹; Sigma), carbenicillin (6 μg ml⁻¹; Sigma), methicillin (16 μg ml⁻¹; Sigma), teicoplanin (0.07 μg ml⁻¹; Sigma), bacitracin (5 μg ml⁻¹; Sigma), nisin (40 μg ml⁻¹; Sigma), LL-37 (500 μg ml⁻¹; Anaspec). Aliquots of 300 μl of antimicrobial stock solutions were added to the last column of a 96-well flat-bottom plate and serially diluted 1:2 into 150 μl of THY broth. For lysozyme treatments, wells were prepared with increasing concentrations of chicken egg white lysozyme (40,000 units mg⁻¹; Sigma) from 2.5 mg ml⁻¹ to 25 mg ml⁻¹ in a 150-μl total volume of THY broth. Mid-exponential-phase bacterial cultures were washed twice in PBS and normalized to 1 × 10⁵ CFU ml⁻¹. Ten microliters of this resuspension was added to each well, and plates were incubated overnight at 37°C. OD₆₀₀ readings were determined using a VersaMax microplate reader (Molecular Diagnostics). The MIC was scored as the lowest antimicrobial concentration with no detectable growth. For assays using LL-37, antimicrobial assays were performed as described above except that PBS was used in place of THY broth. Plates were incubated at 37°C for only 3 h, followed by serial dilution and plating on THY agar. The minimal bactericidal concentration (MBC) was scored as the lowest antimicrobial concentration with no detectable growth.

Bacterial surface charge assay.

Bacteria grown in THY medium were normalized to an OD₆₀₀ of 6.0. Cultures were washed 2 times in 20 mM morpholinepropanesulfonic acid (MOPS) buffer (pH 7.0) and resuspended in 1 ml of the same buffer. Ten microliters of 50-mg ml⁻¹ cytochrome c (Sigma) was added, mixed well, and incubated for 15 min at room temperature. Samples were centrifuged at 13,000 × g for 5 min, supernatants were collected, and the OD₅₃₀ was measured to determine the amount of cytochrome c remaining. Tubes containing no bacteria were used as an input control. The percentage of cytochrome c bound was calculated using the following formula: [1 − (output OD₅₃₀/input OD₅₃₀)] × 100.

RNA isolation and quantitative PCR.

Ten-milliliter aliquots of bacterial cultures were used to purify total RNA as previously described (2). For synthesis of cDNA, 2 μg of total RNA was combined in 20-μl reaction volumes with 0.2 μM reverse primer mix (4 μM mixture of each of the reverse primers listed below), 0.5 mM deoxynucleoside triphosphate mix, 1× reverse transcription buffer, and 200 U of Maxima reverse transcriptase (Fermentas). Reaction mixtures were incubated at 60°C for 30 min, followed by 85°C for 5 min. For qPCRs, 1 μl of cDNA was combined in 20-μl total reaction volumes with 0.2 μM forward and reverse primers and 1× Maxima SYBR green master mix (Fermentas) per the manufacturer's instructions. Primer sequences were as follows: recA (Sin.996), forward, 5′-CTCAGGTGCTGTTGATTTGG-3′, and reverse, 5′-TGCAGAGAGTTTACGCATGG-3′; murA (Sin.1445), forward, 5′-CGTAGTCGTGTATTT-3′, and reverse, 5′-TTGGGTTGTATGGTGAAGCA-3′; oatA (Sin.29), forward, 5′-CTTTTCGGCTTTGTTCTTGC-3′, and reverse, 5′-TCCGCAAAGTATGTCAGCAC-3′; dltA (Sin.295), forward, 5′-CACCGGTATGGTAAGCAGGT-3′, and reverse, 5′-AAACGGAGAGCAAGGTGAAA-3′; pgdA (Sin.822), forward 5′-GATGGTCCAAACCCTGTGAC-3′, and reverse, 5′-ACCCGCTTCCTTGACTTTTT-3′; tagO (Sin.1228), forward, 5′-GGTTCCTATGCCAAGTGGTG-3′, and reverse, 5′-ATTGCCGCTCCAATAACAAC-3′. Reactions were performed in triplicate using a Bio-Rad iCycler under the following cycling conditions: 10 min at 95°C and 40 cycles of 15 s 95°C and 1 min 60°C. Melting curves were performed after completion of each experiment. The relative fold change in gene expression was calculated based on 2^−ΔΔCT as previously described (2).

Isolation and purification of peptidoglycan.

S. iniae strains subcultured in 1 liter of THY at 37°C to early stationary phase were rapidly cooled on ice for 5 min and harvested by centrifugation (9,000 × g, 4°C) for 10 min. The pellet was resuspended in 15 ml of cold water and transferred dropwise into 50 ml of boiling 4% sodium dodecyl sulfate (SDS; Sigma) for 30 min. The SDS-insoluble fraction was collected by centrifugation (9,000 × g, 25°C) for 10 min and washed with 60°C water until the SDS was undetectable by methylene blue assay (30). The cell wall fraction was resuspended in 1 ml of 100 mM Tris-HCl (pH 7.5), 20 mM MgSO₄ containing 10 μg ml⁻¹ DNase I (Qiagen), 250 μg ml⁻¹ RNase A (Qiagen) and incubated for 2 h at 37°C. Calcium chloride was adjusted to a final concentration of 10 mM, and 100 μg trypsin was added to the mixture prior to incubation overnight at 37°C. The samples were boiled in 1% SDS for 5 min to denature trypsin, and the samples were washed clean of SDS as described before. The purified peptidoglycan was lyophilized and stored for later analysis.

Analysis of muropepetide composition.

Muropeptides were analyzed as previously described, with minor variations (49). Five milligrams of lyophilized cell wall preparation was resuspended in 500 μl of 0.1 M Tris-HCl (pH 7) and 20 μl of mutanolysin (5,000 U ml⁻¹; Sigma) and incubated at 37°C overnight. Samples were boiled for 5 min, and the insoluble fraction was separated by centrifugation at 13,000 × g for 2 min. The muropeptide-containing supernatants were lyophilized and resuspended in 5 μl of a 0.2 M disodium 8-amino-1,3,6-naphthalenetrisulfonate (ANTS; Invitrogen) solution in 15% acetic acid (Fisher) and 5 μl of a 1 M sodium cyanoborohydride (Acros) solution in dimethyl sulfoxide. The samples were incubated overnight at 37°C, dried by centrifugal vacuum, and resuspended in 200 μl of loading buffer (62.5 mM Tris-HCl [pH 6.8], 20% glycerol). One-micromole volumes of carbohydrate standard of glucose (Sigma), maltose (Fisher), maltotriose (Acros), maltotetraose (MP Biomedicals), maltopentaose (Sigma), and maltohexaose (MP Biomedicals) were also ANTS labeled and resuspended in 1 ml of loading buffer. Labeled standards were combined to a final concentration of 1 nmol μl⁻¹ each.

Fluorophore-assisted carbohydrate electrophoresis (FACE) was performed in a Hoefer S600 gel apparatus. The 0.5-mm-thick acrylamide gels (18 cm × 16 cm) were composed of a stacking (8% acrylamide, 0.2% bisacrylamide, 0.3 M Tris buffer [pH 8.9]) and running (32% polyacrylamide, 2.4% bisacrylamide, 0.3 M Tris buffer [pH 8.9]) gel component. Fifteen-microliter aliquots of prepared samples were separated by electrophoresis at 4°C for 2.5 h, 35 mA in running buffer (0.192 M glycine, 25 mM Tris [pH 8.9]). The muropeptide bands were visualized by UV exposure (310 nM) and photographed using a Canon Powershot A650IS camera. Gels were analyzed for relative band intensity using Gel-Pro Analyzer software (MediaCybernetics).

Determination of peptidoglycan acetylation.

For quantification of peptidoglycan acetylation, 30-mg aliquots of lyophilized cell wall preparations were resuspended in 1.5 ml 80 mM sodium hydroxide and incubated at 37°C for 3 h with shaking. The samples were ultracentrifuged (100,000 × g, 25°C) for 30 min to separate the insoluble wall fraction. The amount of released sodium acetate in the supernatant was assayed with the acetic acid (AK) kit (Megazyme) using a VersaMax microplate reader (Molecular Diagnostics). Standard curves were prepared from serial dilutions of sodium acetate (Sigma) for quantification.

Statistical analysis.

A statistical analysis for all functional tests was performed by using the two-tailed Student's t test within the StatView analysis software.

RESULTS

CpsY does not affect neutrophil phagosomal maturation.

Because CpsY is critical for intracellular survival in neutrophils (2), we sought to elucidate a mechanism for this protective function. Streptococcus pyogenes has been shown to escape neutrophil phagocytic killing by inhibiting fusion of azurophilic granules with the maturing phagosome (74). To test whether CpsY functions to disrupt neutrophil phagosomal maturation in a similar manner, infected neutrophils were fixed and dual immunostained with antibodies against S. iniae, and either the LAMP-1 marker CD107A or azurophilic granule marker CD63. Laser confocal fluorescence microscopy images revealed that both the WT and the ΔcpsY mutant colocalized with LAMP-1 (Fig. 1A) and azurophilic granule markers (Fig. 1B), implying containment within a mature phagosome. These data suggest that S. iniae does not manipulate phagosomal maturation in neutrophils, as observed for S. pyogenes (74). This also suggests that the protective function imparted by CpsY must occur within a mature neutrophil phagosome.

Fig 1.

S. iniae colocalization with neutrophil phagolysosome markers CD107A and CD63. Infected neutrophils were analyzed by confocal laser scanning fluorescence microscopy for localization of S. iniae WT or ΔcpsY (green) in relation to the human neutrophil LAMP-1 (CD107A; red) (A) or human neutrophil azurophil (primary) granule marker CD63 (red) (B). Shown is a maximum projection of a sequential scan. Diffuse background staining in panel A is medium debris.

Although phagosomal maturation appeared unaltered, CpsY could serve a protective role by regulating factors that function to disrupt the microenvironment within a mature phagosome. To address this possible scenario, a 1:1 coinfection of neutrophils with WT S. iniae and the ΔcpsY mutant was performed to determine if the CpsY protein produced in the WT strain could rescue the mutant strain when inhabiting the same phagosome. Coinfection of neutrophils had no influence on the ΔcpsY mutant, resulting in significant killing compared to WT (P < 0.01) and an extremely low competitive index of 0.06 (Fig. 2). Furthermore, when neutrophil phagocytosis was inhibited by pretreatment with cytochalasin D (CD) or heat inactivation of serum (HIS), growth of the ΔcpsY mutant was restored to levels observed in serum alone (Fig. 2). These survival patterns mirrored that of neutrophils infected individually with either WT S. iniae or the ΔcpsY mutant (2), indicating that CpsY does not function to manipulate the neutrophil phagosome, but rather must regulate factors that provide direct protection to S. iniae from intracellular neutrophil killing mechanisms within a mature phagosome.

Fig 2.

Neutrophil competition assay. Mid-logarithmic cultures of S. iniae WT and ΔcpsY were inoculated at a 1:1 ratio into 1 ml of Dulbecco's modified Eagle's medium with 50% human serum (S) or 50% HIS. Human neutrophils (N) were added at a ratio of 1,000:1 (N:bacteria). If specified, neutrophils were pretreated with CD at 10 μg ml⁻¹ for 30 min to inhibit phagocytosis. Samples were incubated at 37°C for 3 h with gentle rotation, followed by serial dilution and replica plating on THY plates with or without 1,000 μg ml⁻¹ streptomycin for enumeration of bacterial CFU. The CI was determined as described in Materials and Methods. Error bars represent standard errors. *, P < 0.001.

The ΔcpsY mutant has increased lysozyme sensitivity.

Many of the bactericidal compounds contained within a mature neutrophil phagosome target components of the bacterial cell wall. The susceptibility of the ΔcpsY mutant to neutrophil-mediated killing could imply that CpsY regulates factors involved in cell wall physiology. This could manifest as decreased resistance to cell wall-targeting antibiotics or antimicrobial peptides. No differences in the MICs were observed for the antibiotics colistin, gramicidin, carbenicillin, methicillin, teicoplanin, bacitracin, or the antimicrobial peptides nisin and LL-37 (Table 1). However, the ΔcpsY mutant was significantly more sensitive to lysozyme, with a 50% decrease in the MIC compared to WT (Table 1). The increased sensitivity of the ΔcpsY mutant specifically to lysozyme suggested possible chemical alterations in the cell wall (4). In support of this, the ΔcpsY mutant displayed an altered surface charge compared to the WT strain as determined by cytochrome c binding (Fig. 3). Taken together, these data provide evidence for the role of CpsY in regulating factors involved with cell wall physiology.

Table 1.

MICs of tested antibiotics for the two strains

Strain	MIC (mg/ml)
	Colistin	Gramicidin	Carbenecillin	Methecillin	Teichoplanin	Bacitracin	Nisin	LL-37a	Lysozymeb
WT	250	0.0059	0.375	0.5	0.035	2.5	5	31.25	650,000
ΔcpsY	250	0.0059	0.375	0.5	0.035	2.5	5	31.25	300,000c

^aMinimal bactericidal concentration.

^bActive enzymatic units.

^cP = 0.0003 compared to WT.

Fig 3.

Measurement of bacterial cell surface charge. S. iniae WT or ΔcpsY cultures were normalized to an OD₆₀₀ of 6.0, washed twice, and resuspended in 20 mM MOPS buffer with 0.5 mg ml⁻¹ cytochrome c for 5 min. Samples were centrifuged, and the OD₅₄₀ of the supernatant was measured. Data represent the percentage of bound compared to input control. Error bars represent standard errors. *, P < 0.01.

Deletion of cpsY reduces peptidoglycan acetylation.

Lysozyme is an important component of epithelial secretions and phagocytic cells (9, 16, 17), and thus an obstacle necessary for pathogenic species to overcome. Lysozyme cleaves the β-(1,4)-glycosidic bonds between the N-acetylmuramyl (MurNAc) and N-acetylglucosamyl (GlcNAc) residues of peptidoglycan (PG). Chemical modifications to the MurNAc residues of PG that increase lysozyme resistance include both the addition of O-linked acetyl groups by the OatA acetyltransferase (3) and removal of N-linked acetyl groups by PgdA deacetylase (82). Furthermore, the addition of wall teichoic acids and their d-alanylation have been shown to affect susceptibility to many of the antimicrobial peptides contained within neutrophil granules (64). To determine if CpsY influences the expression of genes involved in these processes, we performed qPCR on both mid-logarithmic- and stationary-phase cultures of WT S. iniae and the ΔcpsY mutant. Expression of a putative PG O-acetyltransferase (oatA) was decreased >2-fold in the ΔcpsY mutant relative to WT (Table 2), suggesting that CpsY functions as a transcriptional activator of oatA. No change in expression was observed for genes involved in teichoic acid biosynthesis (tagO), d-alanylation of teichoic acids (dltA), or PG N-deacetylation (pgdA).

Table 2.

qPCR results

Gene	Function	GPa	Fold changeb
murA	MurNAc-l-alanine amidase	ML	2.41 ± 0.13
		ST	2.04 ± 0.01
oatA	O-acetyltransferase	ML	0.43 ± 0.07
		ST	0.39 ± 0.05
dltA	d-Alanine-poly(phosphoribitol) ligase	ML	1.09 ± 0.12
		ST	1.09 ± 0.12
pgdA	GlcNAc deacetylase	ML	1.09 ± 0.15
		ST	1.37 ± 0.70
tagO	UDP-GlcNAc 1-P transferase	ML	1.17 ± 0.18
		ST	0.95 ± 0.09

^aGP, growth phase; ML, mid-logarithmic; ST, stationary.

^bRelative fold change in gene expression for ΔcpsY compared to WT as determined by the ΔΔC_T method.

To determine whether the decreased expression of oatA in the ΔcpsY mutant was accompanied by an actual decrease in PG acetylation, purified PG preparations were treated briefly with 80 mM NaOH to release all O-linked acetyl groups (4). Quantification of released acetyl groups revealed a >20% decrease in the acetylation of PG from the ΔcpsY mutant (300.58 ± 23.08 pmol/mg for WT versus 238.32 ± 10.16 pmol/mg for ΔcpsY; P < 0.05, Student's t test). Thus, the increased susceptibility of the ΔcpsY mutant to lysozyme and intracellular killing by neutrophils may be due to an inability to properly acetylate the cell wall.

The ΔcpsY mutant contains altered muropeptide profiles.

The CpsY ortholog MtaR of S. agalactiae was previously shown to regulate the expression of a diverse gene set (10). In an attempt to gain further insight into the S. iniae CpsY regulon, the microarray data obtained from that study was examined for differential expression of genes with probable effects on cell wall physiology. Relative expression of a gene encoding a putative N-acetylmuramoyl-l-alanine amidase (SAN_0845) was found to be increased over 2-fold compared to WT in that study. Analysis of the S. iniae ortholog (murA) by qPCR revealed the same 2-fold increase in expression for the ΔcpsY mutant relative to WT (Table 2), indicating that CpsY may function as a transcriptional repressor of murA expression.

N-acetylmuramoyl-l-alanine amidases (EC 3.5.1.28) break down PG muropeptides (single disaccharide of GlcNAc and MurNAc with a peptide component) by hydrolyzing the bond between the muramoyl residue of the glycan backbone and the l-amino acid residue of the peptide side chain, a process required for cell division and PG recycling (68). Overproduction of these amidases could result in disruption of the muropeptide unit, resulting in an unstable cell wall and an increased susceptibility to neutrophil bactericidal killing mechanisms. To determine if CpsY influenced the S. iniae muropeptide composition, muropeptides were prepared by mutanolysin treatment of purified PG from WT S. iniae and the ΔcpsY mutant. These samples were then analyzed by FACE (86), which allows for the differentiation of muropeptides based on their electrophoretic mobility profiles. FACE analysis revealed that the relative concentrations of soluble oligomeric muropeptide structures (cross-linked muropeptides) in the ΔcpsY mutant were approximately half that of the WT strain (Table 3, M1 to M5). Furthermore, the monomeric structures (individual monosaccharides of MurNAc or MurNAc plus peptides) were increased (Table 3, M7 and M9). These results demonstrated that the ΔcpsY mutant has an altered PG muropeptide profile suggestive of a weakened cell wall; however, this is likely an additive effect in combination with other processes (i.e., decreased acetylation), because overexpressing murA alone in WT S. iniae did not completely alter the muropeptide profile to that of the ΔcpsY mutant (data not shown).

Table 3.

Compositions of muropeptides

Muropeptide	Compounda	% muropeptide composition inb:		P valuec
		WT	ΔcpsY
M1	Oligomeric MurNAc-peptide	6.61 ± 1.28	3.22 ± 0.99	0.20
M2	Oligomeric MurNAc-peptide	3.29 ± 0.42	2.53 ± 1.11	0.57
M3	Oligomeric MurNAc-peptide	3.94 ± 0.37	1.64 ± 0.69	0.07
M4	Oligomeric MurNAc-peptide	15.47 ± 3.62	6.97 ± 2.75	0.22
M5	Oligomeric MurNAc-peptide	7.47 ± 0.98	3.92 ± 1.35	0.20
M6	Oligomeric MurNAc-peptide	5.76 ± 2.78	7.15 ± 5.98	0.86
M7	MurNAc-tripeptide	39.71 ± 1.24	47.66 ± 3.17	0.06
M8	GlcNAc	2.47 ± 0.68	2.31 ± 1.27	0.94
M9	MurNAc	15.29 ± 3.19	24.59 ± 2.61	0.19

^aMuropeptide compound, based on electrophoresis profile relative to known carbohydrate standards (86).

^bThe muropeptide percent composition (mean ± standard error) was determined by FACE analysis.

^cBased on Student's t test.

Analysis of CpsY-regulated promoters.

Previous work has shown that certain genes under transcriptional control by streptococcal orthologs of CpsY contain a highly conserved regulatory element termed the Met box within their promoter region (40, 73). The presence of a Met box in the promoter region of a CpsY-regulated methionine ABC transport system (AtmBDE) suggests that this same element facilitates CpsY-dependent regulation in S. iniae as well (2). We sought to determine if transcriptional regulation of oatA or murA by CpsY is through this conserved element. The gene encoding OatA is predicted to be the third gene of an operon (Fig. 4A). Whether murA is part of an operon is unknown, but the predicted promoter is immediately upstream of murA (Fig. 4A). Analysis of the predicted promoter regions revealed the presence of a Met box within the murA promoter, but not the oatA operon promoter (Fig. 4B). Additionally, although the promoters of both murA and atmB contain the canonical Met box, a major structural difference exists for those regions. Twenty-two nucleotides separate the Met box and −35 site in the atmB promoter, whereas those elements are separated by 96 nucleotides in the murA promoter (Fig. 4B). This difference may explain why CpsY transcriptionally activates expression of atmB (2) while repressing murA (Table 2). The absence of a Met box for oatA suggests that the effect of CpsY on the expression of oatA may be indirect or occur in combination with additional factors.

Fig 4.

Promoter alignment for CpsY-regulated genes. (A) Genetic locations of AtmBDE (Sin. 1702 to 1699), MurA (Sin. 1445), and OatA (Sin. 29). (B) Predicted promoter sequences obtained from GenBank were aligned and plotted using ClustalX. The predicted −10 and −35 sites are underlined, the translational start codon is italicized, and predicted Met boxes are shaded.

DISCUSSION

Neutrophil phagocytosis of an opsonized pathogen is an intricate process whereby sequential maturation of the phagosome generates a highly bactericidal microenvironment (47, 71). Pathogens within the nascent phagosome are exposed to a burst of reactive oxygen species subsequent to assembly of the NADPH oxidase. Phagosomal maturation ensues, with successive fusion of preformed gelatinase, specific, and azurophilic granules that contain an array of bactericidal compounds (71).

Engulfed microbes endure a multitargeted attack within a mature phagosome. Transporters such as the natural resistance-associated macrophage protein 1 (Nramp1) (35) and chelating agents like neutrophil gelatinase-associated lipocalin (NGAL) (37) and lactoferrin (58) deprive microorganisms of vital metals such as iron, manganese, and zinc. Bacterial lipid membranes are the target of cytotoxic pore-forming antimicrobial peptides, such as cathelicidins (67) and defensins (26) with bactericidal activities against both Gram-negative and Gram-positive bacteria. Furthermore, myloperoxidase can react with hydrogen peroxide to generate hypochlorous acid (HOCl), tyrosine radicals, and reactive nitrogen intermediates, all of which can attack the surface membranes of microorganisms (38). Engulfed pathogens are also exposed to serprocidins (serine proteases with microbicidal activity) like proteinase-3, cathepsin G, and elastase (13, 69, 72), as well as lysozyme, which has both muramidase-dependent and -independent antimicrobial activities (57).

Bacterial pathogens employ 3 characterized strategies for intracellular survival in neutrophils: inhibition of phagosomal maturation, survival within a mature phagosome, or escape into the cytoplasm (79). S. pyogenes evades intracellular neutrophil killing by inhibiting phagosomal maturation through an uncharacterized mechanism involving M protein (74). In contrast, we demonstrated that phagosomal maturation is unaltered in S. iniae-infected neutrophils, as determined by colocalization with azurophilic granules. Thus, the high degree of attenuation observed for the ΔcpsY mutant upon phagocytosis must be due to the dysregulation of specific factors required for survival within a mature phagosome. These CpsY-regulated factors must be acting directly on the bacteria, rather than manipulating the phagosomal microenvironment, based on the inability of WT S. iniae to rescue the ΔcpsY mutant during coinfection of neutrophils.

Protection from the bactericidal components encountered within a mature neutrophil phagosome begins in part with sensory input from a variety of two-component systems (TCS), resulting in the activation of cell stress response and wall modification programs (42, 48, 63, 66, 75, 80). These systems can activate enzymes that allow the bacteria to decorate both the cell membrane and cell wall with various chemical modifications. For example, lipid lysinylation of the cellular membrane by MprF (23, 44, 61, 70, 77) and d-alanylation of wall teichoic acids by the DltABC locus (15, 24, 39, 41, 43, 64) provide protection from cationic antimicrobial peptides. Resistance to lysozyme occurs through PG acetylation of MurNAc residues by the OatA O-acetyltransferase found in Gram-positive organisms (3, 4, 19, 34, 80) or the PatA/PatB system in Gram-negative organisms (56). Additionally, the PgdA N-deacetylase mediates lysozyme resistance through deacetylation of PG GluNAc residues (7, 54, 81–83). The significant increase in lysozyme sensitivity of the ΔcpsY mutant without a concomitant change in sensitivity to β-lactam antibiotics or antimicrobial peptides tested suggests possible specific chemical modifications to PG, such as O-acetylation or N-deacetylation. The decrease in oatA expression for the ΔcpsY mutant with a concomitant reduction in total PG acetylation suggests that susceptibility to neutrophil-mediated killing may be due to the inability to properly acetylate the cell wall.

The OatA O-acetyltransferase was originally identified in pathogenic staphylococci (3, 4), and homologs have been described for Streptococcus pneumoniae (19), Enterococcus faecalis (31), and Lactococcus lactis (80). OatA is a membrane-spanning protein thought to utilize cytoplasmic acetyl coenzyme A as a donor for the translocation of acetate to the C-6 hydroxyl group of MurNAc residues in the murein sacculus (46). In L. lactis the CesSR TCS responds to lysozyme-induced cell envelope stress by activation of OatA through the intermediate SpxB regulator (80). CesSR homologs have also been shown to respond to cell envelope stress in S. pneumoniae (22, 28), S. mutans (75), Staphylococcus aureus (45), Bacillus licheniformis (84) and Bacillus subtilis (36). S. iniae also contains a CesSR homolog; however, its functionality in the cell envelope stress response is unknown. Whether the reduced PG acetylation observed for the ΔcpsY mutant is influenced by the CesSR stress response pathway has yet to be determined. The observed reduction in PG acetylation of the ΔcpsY mutant could alternatively represent an inability to accumulate acetylated PG through exponential growth into stationary phase rather than an effect of a specific stress response (46, 62). Attempts to complement the ΔcpsY mutant through overexpression of oatA were unsuccessful, resulting in slight growth retardation and continued susceptibility to neutrophil-mediated killing in whole blood (data not shown). Similar observations on growth have been made for PG hyperacetylation in L. lactis (80) and Lactobacillus planetarum (5), suggesting that tight regulation of the O-acetyltransferase is necessary for optimal growth.

The observed decrease in PG acetylation for the ΔcpsY mutant could result in an increased sensitivity to the cell's own autolysins (6, 60). These PG hydrolases encompass several distinct enzymes, including the N-acetylmuramidases, N-acetylglucosaminidases, N-acetylmuramyl-l-alanine amidases, and lytic transglycosylases, all of which perform specific enzymatic functions necessary for cell wall growth, turnover, and separation during cellular division (68). The strict spatial and temporal control of these enzymes is critical for prevention of unregulated PG hydrolysis (78), and PG O-acetylation has been shown to inhibit certain autolysins (6, 60). As shown here, CpsY acts as a transcriptional repressor of a putative N-acetylmuramoyl-d-alanine amidase (MurA), presumably through interactions with the conserved Met box in the murA promoter. N-Acetylmuramoyl-l-alanine amidases play an important role in separation during cell division (14, 27, 32, 46, 65) and can be potent autolysins when overexpressed (32, 33). The increased expression of murA along with decreased PG acetylation could render the ΔcpsY mutant susceptible to detrimental restructuring of the murein sacculus, thereby increasing the sensitivity to many of the bactericidal factors contained within a neutrophil phagosome. FACE analysis revealed a reduction in the proportion of oligomeric muropeptide structures (cross-linked soluble muropeptides) for the ΔcpsY mutant with a concomitant increase in single muropeptides and free MurNAc and GluNAc residues, indicative of an altered cell wall structure. Overexpression of MurA alone in WT S. iniae did not produce the same muropeptide profile observed for the ΔcpsY mutant and resulted in little difference from WT for growth or susceptibility to killing in neutrophils in whole blood (data not shown). Thus, simple overexpression of MurA, perhaps without an accompanying decrease in PG acetylation or other CpsY-dependent effects, does not explain the susceptibility to neutrophil-mediated killing observed for the ΔcpsY mutant.

CpsY is a complex transcriptional regulator, and the high degree of conservation is indicative of its critical importance for basic processes of all prokaryotic microorganisms. We demonstrate here that mutation of cpsY in S. iniae appears to alter properties of the cell wall, which may explain the increased susceptibility of the ΔcpsY mutant to neutrophil-mediated killing mechanisms. The inability to ectopically complement the ΔcpsY mutation likely reflects its tight regulatory function and vital role during streptococcal infection. Many different cellular pathways converge on PG biosynthesis and cell wall modification, and mutants in components of these pathways often share similar virulence phenotypes. For example, S. iniae mutants of phosphoglucomutase (PgmA) (12) and S. mutans mutants of phosphoglucosamine mutase (GlmM) (85) also display similar increases in lysozyme sensitivity, susceptibility to neutrophil-mediated killing, and virulence attenuation in vivo. The common attenuation observed upon mutation within these converging pathways suggests that combinational therapies targeting these basic processes would be a powerful treatment strategy. Continued studies into the regulatory function of CpsY using S. iniae as a model pathogen should highlight critical streptococcal adaptations that occur during the course of an infection, providing necessary information about invasive streptococcal disease.