Abstract
Bacterial cell division is driven by a tubulin homolog FtsZ, which assembles into the Z-ring structure leading to the recruitment of the cell division machinery. In ovoid-shaped Gram-positive bacteria, such as streptococci, MapZ guides Z-ring positioning at cell equators through an, as yet, unknown mechanism. The cell wall of the important dental pathogen Streptococcus mutans is composed of peptidoglycan decorated with Serotype c Carbohydrates (SCCs). Here, we show that an immature form of SCC, lacking the recently identified glycerol phosphate (GroP) modification, coordinates Z-ring positioning. Pulldown assays using S. mutans cell wall combined with binding affinity analysis identified the major cell separation autolysin, AtlA, as an SCC binding protein. Importantly, AtlA binding to mature SCC is attenuated due to GroP modification. Using fluorescently-labeled AtlA, we mapped SCC distribution on the streptococcal surface to reveal that GroP-deficient immature SCCs are exclusively present at the cell poles and equators. Moreover, the equatorial GroP-deficient SCCs co-localize with MapZ throughout the S. mutans cell cycle. Consequently, in GroP-deficient mutant bacteria, proper AtlA localization is abrogated resulting in dysregulated cellular autolysis. In addition, these mutants display morphological abnormalities associated with MapZ mislocalization leading to Z-ring misplacement. Altogether, our data support a model in which GroP-deficient immature SCCs spatially coordinate the localization of AtlA and MapZ. This mechanism ensures cell separation by AtlA at poles and Z-ring alignment with the cell equator.
Introduction
Bacterial cells come in a variety of shapes. The specific bacterial shapes are imposed by their cell wall, which surrounds the cytoplasmic membrane. The main structural component of the cell wall is peptidoglycan, which is composed of glycan strands that are cross-linked by penta-peptides. During cell division, new peptidoglycan is synthesized and inserted into the existing cell wall by the coordinated action of enzymes catalyzing peptidoglycan hydrolysis and synthesis. This process is tightly controlled at both the spatial and temporal level to prevent the loss of cell wall integrity and ultimately guarantee the correct cell morphology.
According to the current model, the morphogenesis of ovoid-shaped Gram-positive bacteria, such as streptococci, enterococci and lactococci, arises from a combination of septal and lateral peptidoglycan synthesis, which is coordinated by multiprotein complexes called the divisome and elongasome, respectively 1-3. Cell division is initiated by the recruitment and polymerization of FtsZ to form a structure called the Z-ring at mid-cell marked by a microscopically visible “wall band” or “equatorial ring” 1,4,5. Misplacement of FtsZ leads to severe morphological abnormalities. The Z-ring serves as a scaffold for other components of the cell division machinery, including peptidoglycan polymerases and hydrolases 3,5, that start cell division by synthesizing a small septal ingrowth below the equatorial ring 1. Early in division, new equatorial rings appear in the daughter cells, presumably due to splitting of the parental equatorial ring 1. During the elongation phase, the rings gradually migrate toward the equators of the daughter cells powered by two processes — splitting of the septal ingrowth and synthesis of the lateral wall 1. When the daughter cells have reached the size of the parental cell, and the equatorial ring approaches the mid-cell region of the nascent daughter cell, elongation is halted. At the same time, synthesis of the septal wall rapidly resumes, followed by final splitting of the complete septum by peptidoglycan hydrolases, or so-called autolysins, to allow the proper separation of the daughter cells 1,4,5.
It remains unclear what cues in the cell wall direct the recruitment of the cell separation autolysins. Furthermore, it is unknown how FtsZ is targeted to mid-cell. Currently, it is assumed that the correct placement of the Z-ring depends on the chromosomal origin of replication 6 and the FtsZ-binding protein MapZ protein MapZ 7,8. MapZ forms a stable protein ring that co-migrates with the equatorial ring 7,8, facilitating the alignment of the Z-ring perpendicular to the long axis of the cell 6. In Streptococcus pneumoniae, the MapZ-ring acts as a continuous guide for the orderly migration of FtsZ from the parental septum to the equatorial rings of daughter cells throughout the cell cycle 9. In contrast, Streptococcus mutans MapZ promotes FtsZ movement to the equators of daughter cells at a later stage in division 10. MapZ localization and assembly into the ring structure are reported to depend on the direct interaction of the MapZ extracellular domain with an unknown cell wall component residing in the equatorial ring 7,11.
The cell wall of Gram-positive bacteria contains characteristic anionic glycopolymers covalently attached to peptidoglycan. The best-studied class of cell wall glycopolymers is the canonical poly(glycerol-phosphate) and poly(ribitol-phosphate) wall teichoic acids (WTAs) expressed by Bacillus subtilis and Staphylococcus aureus, respectively 12. In these species, WTA-deficient mutants exhibit cell shape abnormalities, defects in the separation of daughter cells, and increased autolysis 13-17, indicating a functional connection between WTA and the cell division machinery. Many streptococci, including the important human dental pathogen S. mutans, lack canonical WTAs. Instead, they express rhamnose (Rha)-containing polysaccharides with a conserved repeating →3)α-Rha(1→2)α-Rha(1→ disaccharide backbone modified with species-specific and serotype-specific glycosyl side-chains. S. mutans strains are classified into four serotypes based on variations in the glycosyl side-chains, with serotype c being the most common in the oral cavity 18,19. The polysaccharide in S. mutans serotype c is referred to as serotype c-specific carbohydrate (SCC) and contains α-glucose (Glc) side-chains attached to the 2-position of the α-1,3 linked Rha 20. The homologous polysaccharide in Streptococcus pyogenes (Group A Streptococcus or GAS), referred to as the Lancefield group A carbohydrate (GAC), carries β-N-acetylglucosamine (GlcNAc) side-chain modifications attached to the 3-position of the α-1,2 linked Rha 21,22. There is substantial evidence that SCC and GAC are critical for cell division of streptococci 23-27. We recently demonstrated that SCC and GAC are also negatively charged polysaccharides, similar to WTAs, through decoration with glycerol phosphate (GroP) moieties 28. NMR analysis of GAC revealed that GroP is attached to the GlcNAc side-chains of GAC at the C6 hydroxyl group 28.
In this study, we link GroP modification of SCC to spatial regulation of streptococcal cell division. We show that structurally-diverse SCCs display a specific distribution on the S. mutans cell surface with cell equators and poles being populated by ‘immature SCCs’, which are deficient in the GroP modification. These immature SCCs inform the proper positioning of MapZ and the major cell separation autolysin AtlA. Thus, the presence of GroP-modified SCC in the streptococcal cell wall provides an exclusion strategy for critical cell division proteins involved in the first and the final stages of streptococcal cell division.
Results
GroP is attached to the Glc side-chains of SCC
We have previously shown that GroP attachment to SCC and GAC is catalyzed by a dedicated GroP transferase encoded by sccH/gacH 28. This gene is located in the 12-gene loci, sccABCDEFGHMNPQ (Fig. 1a), and gacABCDEFGHIJKL 29, which encode the biosynthesis machinery for SCC and GAC, respectively. Because in GAS, GacH is required to transfer GroP to the GlcNAc side-chains of GAC 28, we suggested that SccH similarly modifies the Glc side-chains of SCC with GroP in S. mutans. To test this hypothesis, we generated S. mutans c serotype strains that were devoid of the Glc side-chains. In GAS, the GtrB-type glycosyltransferase, GacI, is critical for GAC GlcNAc side-chain modification through the formation of the donor molecule GlcNAc-phosphate-undecaprenol 30. The SCC gene cluster (Fig. 1a) contains two putative GtrB-type glycosyltransferases encoded by sccN and sccP. To investigate the function of SccN and SccP in the synthesis of the Glc donor for side-chain addition to SCC, we deleted sccN and sccP in S. mutans serotype c strain Xc, creating ΔsccN, ΔsccP, and the double mutant ΔsccNΔsccP. The glycosyl composition of purified SCCs exhibited a significantly reduced amount of Glc in ΔsccN and ΔsccNΔsccP (Fig. 1b). In contrast, the deletion of sccP alone did not affect Glc levels. The Glc content of SCC was restored in ΔsccN by complementation with sccN on an expression plasmid (ΔsccN:psccN, Fig. 1b). These data strongly support a major role for SccN in Glc side-chain formation of SCC. Interestingly, the Glc content of SCC in ΔsccNΔsccP was lower than in ΔsccN (Fig. 1b), suggesting that sccP might play a minor role in providing a Glc donor for modification of SCC with the side-chains (Supplementary Fig. 1a). Furthermore, we found that the glycerol and phosphate content in the polysaccharide isolated from ΔsccN was significantly reduced (Fig. 1b), similar to the ΔsccH mutant (Fig. 1b). We have previously provided conclusive evidence that glycerol and phosphate detected in this analysis are, in fact, GroP 28. The deficiency of glycerol and phosphate in the ΔsccN mutant was reversed by complementation with sccN, supporting the conclusion that the Glc-side chains of SCC are further modified with GroP by SccH (Supplementary Fig. 1a, b and c).
GroP modification controls the self-aggregation and morphology of S. mutans
We observed that planktonic ΔsccH and ΔsccN have a strong tendency to spontaneously sediment after overnight growth, as compared to the wild type (WT) strain and the complemented strains, ΔsccH:psccH and ΔsccN:psccN, which remained in suspension (Fig. 2a). Microscopic analysis of bacteria revealed that ΔsccH and ΔsccN, but not the WT strain, ΔsccH:psccH and ΔsccN:psccN, formed the typical short chains that clump together (Fig. 2b). Furthermore, the bacterial aggregates of ΔsccH and ΔsccN were not dispersed when DNAse was added to the growth medium (Supplementary Fig. 2), suggesting that this bacterial behavior is due to cell-cell interactions.
Analysis of WT, ΔsccH, and ΔsccN by scanning electron microscopy (SEM) and differential interference contrast (DIC) microscopy revealed that the mutant cells have severe cell division defects. The WT bacteria displayed the characteristic oval shape with the average cell length of 0.88±0.11 µm in the exponential growth phase (Fig. 2c, d, and e and Supplementary Table 3). In contrast, the majority of the ΔsccH and ΔsccN cells were significantly shorter than the WT cells with the average length of 0.74±0.14 µm and 0.78±0.18 µm, respectively (Fig. 2e and Supplementary Table 3). Additionally, the distribution in the length of the individual ΔsccH and ΔsccN cells differed significantly compared to WT (Supplementary Fig. 3); 21% of the ΔsccH cells (n = 86) and 14% of the ΔsccN cells (n = 117) were abnormally small, while only 2% of the WT cells displayed the minimal cell length (Supplementary Table 3). The small cells of ΔsccH and ΔsccN were frequently paired with larger cells (Fig. 2c and d), implying that these cells resulted from asymmetrically cell division, giving rise to daughter cells with unequal sizes. Finally, cells with the orientation of the division plane not perpendicular to the long axis of the cell were also observed in the ΔsccH and ΔsccN mutants (Fig. 2c). The morphological phenotypes were restored to WT in ΔsccH:psccH and ΔsccN:psccN (Fig. 2c, e and Supplementary Table 3). These phenotypes were also correlated with a significant decrease in cell viability of ΔsccH determined by colony-forming unit count, in comparison to WT and ΔsccH:psccH (Supplementary Fig. 4). We thus concluded that either GroP or the epitope presented by the Glc side-chains modified with GroP are required for proper morphogenesis of S. mutans.
To dissect the structural requirements underlying these morphological processes, we replaced the SCC Glc side-chains with GlcNAc in S. mutans by plasmid-based expression of gacHIJKL 30 in ΔsccN creating ΔsccN:pgacHIJKL (Supplementary Fig.1d and e). These genes (Supplementary Fig. 1e) are required for the formation and addition of the GAC GlcNAc side-chains and GroP in GAS 30. Additionally, as a negative control, we expressed gacHI*JKL in the ΔsccN genetic background creating ΔsccN:pgacHI*JKL. This strain does not synthesize the GlcNAc donor for side-chain addition since it carries a stop codon in gacI. The cell wall polysaccharide purified from ΔsccN:pgacHIJKL showed increased levels of GlcNAc and GroP (Fig. 2f), whereas GlcNAc and GroP were not restored by expression of gacHI*JKL in ΔsccN (Fig. 2f). The morphological phenotypes of ΔsccN were only reversed to WT by expression of gacHIJKL but not gacHI*JKL (Fig. 2a, b, c, e, Supplementary Fig.3 and Supplementary Table 3). These observations indicate that the underlying molecular mechanisms for clumping behavior and the morphological abnormalities are GroP-dependent and independent of the specific glycosyl side chain. The simple explanation for the self-aggregation of bacteria is that ΔsccH and ΔsccN lack a negative surface charge provided by GroP, which normally contributes to electrostatic repulsion between the WT cells.
GroP modification protects S. mutans from autolysis
It has been reported that the WTA-deficient mutant of S. aureus show increased fragility and autolysis due to mislocalization and increased abundance of the major cell division autolysin, Atl, on the bacterial surface 15. Hence, to understand whether and how GroP and the Glc side-chains in SCC affect S. mutans autolysis, we compared autolysis of the WT, ΔsccH, ΔsccN and the complemented strains ΔsccH:psccH, ΔsccN:psccN, ΔsccN:pgacHIJKL and ΔsccN:pgacHI*JKL by analyzing the change in OD600 in liquid cultures after the addition of a mild detergent Triton-X100. While S. mutans WT is relatively resistant to detergent-induced lysis (Fig. 3a), ΔsccH and ΔsccN are sensitive to the autolytic effect of Triton X-100. Cellular lysis was more pronounced in ΔsccN, than in ΔsccH, resulting in significant loss of turbidity of the mutant suspension after 2 hours (Fig. 3a). The phenotypes of ΔsccH and ΔsccN were reversed to WT in ΔsccH:psccH and ΔsccN:psccN. Furthermore, expression of gacHIJKL, but not gacHI*JKL, in the ΔsccN mutant restored the resistance of bacteria to autolysis. These data indicate the importance of GroP and side-chain modifications of SCC in protecting S. mutans from adventitious cellular lysis by an unknown autolytic enzyme.
AtlA promotes autolysis of the mutants lacking GroP and side-chain modifications
A possible mechanistic explanation for enhanced autolysis of ΔsccH and ΔsccN could be that the absence of GroP and Glc side-chain modifications in SCC causes a dysregulated localization of an unknown autolysin. To identify autolytic enzymes that interact with SCCs, we stripped the cell surface-associated proteins from S. mutans WT cells using 8 M urea, re-folded the released proteins by dialysis, and used the re-folded proteins for pulldown experiments with the protein-deficient cell wall material purified from S. mutans. Two major proteins, isolated by the pulldown approach, were identified as AtlA and SmaA by LC-MS/MS analysis (Fig. 3b). One of these, AtlA, is the major autolysin involved in the separation of daughter cells after division 31-34. The protein contains an N-terminal putative cell wall-binding domain, which consists of six Bsp repeats (PF08481) and a C-terminal family GH25 catalytic domain (PF01183) (Fig. 3c). SmaA has two Bsp repeats and three bacterial Src homology 3 (SH3) domains (Fig. 3c). Although SmaA has been implicated in peptidoglycan hydrolysis 35, the protein lacks a putative catalytic domain.
To confirm the identities of the proteins, we generated ΔatlA and ΔsmaA deletion mutants. Using a similar approach as described above, surface proteins were stripped from the ΔatlA and ΔsmaA bacteria and used in pulldown experiments. In agreement with the results of protein identification, the bands corresponding to AtlA and SmaA were absent in ΔatlA and ΔsmaA, respectively (Fig. 3b). To investigate the contribution of AtlA and SmaA in Triton X-100-induced autolysis of WT, ΔsccH, and ΔsccN, we deleted atlA in the ΔsccH and ΔsccN backgrounds, and smaA in the ΔatlA and ΔsccN backgrounds, generating the ΔsmaAΔatlA, ΔsccHΔatlA, ΔsccNΔatlA, and ΔsccNΔsmaA mutants. As expected, the ΔatlA, ΔsmaA, and ΔsmaAΔatlA mutants are relatively resistant to detergent-stimulated lysis (Fig. 3a, Supplementary Fig. 5). The ΔsccNΔsmaA mutant displayed increased cellular lysis, similar to ΔsccN, indicating that SmaA is not involved in detergent-induced autolysis of ΔsccN (Supplementary Fig. 5b). In contrast, the deletion of atlA in the ΔsccH and ΔsccN backgrounds restored the bacterial resistance to the autolytic effect of Triton X-100 (Fig. 3a). These results clearly show that autolysis of ΔsccH and ΔsccN is AtlA-mediated.
Next, we investigated whether the deletion of SmaA or AtlA affected morphology and self-aggregation of S. mutans. The ΔsmaA cells remained in suspension after overnight growth (Fig. 3d), and showed WT cell morphology, cell separation, and cell length distribution (Fig. 3e, f, Supplementary Fig. 3 and Supplementary Table 3). The ΔatlA cells grew in long chains (Fig. 3e) that settle at the bottom of the tube (Fig. 3d), which is in line with the published observation 33. However, in contrast to ΔsccH and ΔsccN, the individual ΔatlA cells displayed normal dimensions, and the long chains of ΔatlA did not self-aggregate (Fig. 3e, f, Supplementary Fig. 3, and Supplementary Table 3). Finally, we examined the phenotype of the ΔsccHΔatlA mutant to understand more about the link between AtlA and GroP modification of SCC. The phenotype of ΔsccHΔatlA was a combination of the phenotypes of single mutants. Notably, similar to ΔatlA, the double mutant displayed a chaining phenotype, and like ΔsccH, the long chains of the double mutant formed clumps (Fig. 3e), supporting the hypothesis that self-aggregation of ΔsccHΔatlA is due to loss of electrostatic repulsion between the cells. Furthermore, SEM and DIC analyses of ΔsccHΔatlA revealed a variety of aberrant cell shapes and size that contrast with the morphology of WT and ΔatlA, but similar to ΔsccH. (Fig. 3e, f, Supplementary Fig. 3 and Supplementary Table 3). Altogether, these data indicate that, although autolysis of ΔsccH and ΔsccN requires AtlA, the cell shape abnormalities observed in ΔsccH and ΔsccN are not linked to either SmaA or AtlA.
GroP modification coordinates the localization of AtlA at the cell pole and mid-cell
Considering the established function of AtlA in the autolysis of ΔsccH and ΔsccN, we hypothesized that the enzyme is either mislocalized or over-expressed in these mutants. To examine the expression of AtlA, the protein was extracted from the cell surface of the WT, ΔsccH, and ΔsccN bacteria. Western blot analysis with anti-AtlA antibodies revealed no significant differences in the amount of AtlA recovered from S. mutans WT or the mutants (Supplementary Fig. 6).
Next, we monitored the localization of AtlA on the cell surface of WT, ΔsccH, and ΔsccN by immunofluorescent microscopy using anti-AtlA antibodies and fluorescent secondary antibodies (Fig. 3g). The autolysin localized to the cell poles and the mid-cell of the WT strain. Secondary antibodies alone showed no specific binding to S. mutans (Supplementary Fig. 7a). In the case of ΔsccH and ΔsccN, the autolysin was evenly distributed over the cell surface, and showed no distinct surface localization pattern (Fig. 3g).
To provide a more detailed picture of the bacterial regions targeted by AtlA, we fused the N-terminal Bsp repeat domain of AtlA with a green fluorescent protein (GFP) 36, generating AtlA-GFP (Fig. 3c). The fusion protein was added exogenously to exponentially growing S. mutans WT, ΔsccH, ΔsccN, ΔsccH:psccH and ΔsccN:psccN cells. Fluorescence microscopy imaging indicated that in newborn WT cells, AtlA-GFP predominantly targeted poles and mid-cell zones (Fig. 3h). Surprisingly, in the cells that were beginning to elongate, the sites labeled by incubation with AtlA-GFP split as a pair of rings (Fig. 3h). Splitting of the mid-cell sites was observed in 47% of the cells (64/135). The mid-cell localization of AtlA-GFP in newborn cells and the duplicated AtlA-GFP localization signal in elongating cells correlate with the position of the equatorial ring in ovococci 4,37. In contrast, AtlA-GFP added to ΔsccH and ΔsccN was uniformly distributed along the bacterial cell surface (Fig. 3h). The complemented strains ΔsccH:psccH and ΔsccN:psccN demonstrated localization of AtlA-GFP similar to the WT strain (Fig. 3h). GFP alone failed to bind to the bacteria (Supplementary Fig. 7b). Together our findings provide three important insights: i) the N-terminal Bsp repeat domain of AtlA is sufficient for proper binding of AtlA to the bacterial surface; ii) the Bsp repeat domain recognizes the specific cell wall component incorporated in cell poles and the regions corresponding to equatorial rings; iii) the absence of GroP and side-chain decorations on SCC correlate with mislocalized binding of AtlA across the entire surface of S. mutans.
AtlA binds to SCC
To identify the cell surface structure targeted by AtlA-GFP, we examined the binding of the protein fusion to sacculi purified from S. mutans WT, ΔsccH and ΔsccH:psccH. The sacculi were free of LTA, proteins, lipids and nucleic acids 30. Fluorescence microscopy imaging revealed that AtlA-GFP associated with the sacculi derived from these cells with patterns very similar to that observed with intact S. mutans cells (Fig. 3i). AtlA-GFP was found primarily at the poles and mid-cell sites of WT and ΔsccH:psccH sacculi, but was distributed evenly along the surface of ΔsccH sacculi. Furthermore, the splitting of AtlA-GFP-labeled mid-cell sites in WT and the complemented cells was observed. Splitting was detected in 50% of WT cells (63/125). GFP alone was unable to attach to the sacculi (Supplementary Fig. 7c). This observation confirmed that the N-terminal domain of AtlA recognizes a cell wall component, either SCC or peptidoglycan, associated with the cell equators and poles.
Notably, secreted proteins consisting of multiple Bsp repeats are widespread in Firmicutes, with most bacteria belonging to the Streptococcus genus (Supplementary Fig. 8). Intriguingly, there is an obvious correlation between the co-occurrence of the genes encoding proteins with Bsp repeats and Rha-containing cell wall polysaccharides in these bacteria (Supplementary Table 4). In fact, in many streptococci, a gene encoding a putative autolysin with Bsp repeats is situated immediately downstream of the gene loci involved in the biosynthesis of the Rha-containing polysaccharide, raising the possibility that the Bsp repeat domain binds Rha-containing cell wall polysaccharides. Thus, to examine the role of SCCs in the recruitment of AtlA, we applied a co-sedimentation assay that exploits the property of AtlA-GFP to associate with the cell wall material purified from WT S. mutans, ΔsccH and ΔsccH:psccH. In agreement with the fluorescent microscopy experiments, we observed a very strong binding of the fusion protein to WT S. mutans, ΔsccH and ΔsccH:psccH cell walls (Fig. 4a). GFP alone did not attach to cell walls purified from S. mutans WT (Fig. 4b). Next, we chemically cleaved the polysaccharides from peptidoglycan using mild acid hydrolysis. SCCs were efficiently released from the cell walls in these conditions (Supplementary Fig. 9a). We observed a significant reduction in AtlA-GFP binding to the SCC-depleted cell wall (Fig. 4a), supporting the role of SCCs in targeting of AtlA to the equatorial rings and the poles of the WT bacteria. To further investigate the participation of SCCs in cellular localization of AtlA, we constructed a SCC-deficient mutant by in-frame deletion of the rgpG gene whose product catalyzes the first step in SCC biosynthesis 38. The ΔrgpG mutant was devoid of polyrhamnose polysaccharide (<0.5% WT level), and in agreement with the published data 39,40, demonstrated self-aggregation and aberrant morphology (Supplementary Fig. 9a and b). Applying AtlA-GFP to the intact ΔrgpG bacteria (Supplementary Fig. 9c) and to the cell wall purified from ΔrgpG (Fig. 4b), no evidence of binding was observed in either assay. These results strongly indicate that SCC is a binding receptor of AtlA.
AtlA binds to SCC via the polyrhamnose backbone
The ability of AtlA to target distinct sites on the S. mutans cell might be explained by binding of the autolysin to a specific form of SCC present in the equatorial rings and the poles. Our observation that AtlA-GFP is uniformly distributed over the cell surface of the side-chain-deficient mutant, ΔsccN, suggests that the N-terminal domain of AtlA binds to polyrhamnose regions on SCC. To determine if AtlA specifically binds the polyrhamnose backbone of SCC we compared the binding of AtlA-GFP to intact cells, or purified cell walls, of bacterial strains that express cell wall polysaccharides containing this backbone feature (GAS and Streptococcus equi) 29 with other bacterial strains that express Rha-containing cell wall polysaccharides lacking this structure (Group B Streptococcus or GBS, and Enterococcus faecalis) (Supplementary Fig. 10a, b, c, d and e). No significant binding was observed to the cell walls or intact cells of GBS and E. faecalis (Fig. 4b and Supplementary Fig. 10f). In contrast, as compared to GFP control, AtlA-GFP strongly binds to the cell wall material purified from GAS (Fig. 4b) and the intact GAS and S. equi cells (Supplementary Fig. 10f), suggesting that the polyrhamnose backbone of GAC, SCC and the S. equi cell wall polysaccharide is recognized by AtlA.
To gather additional evidence that AtlA interacts with the polyrhamnose backbone, we took advantage of a previously developed heterologous expression model in E. coli. Specifically, complementation of the SCC polyrhamnose biosynthetic genes sccABCDEFG in E. coli results in the decoration of lipopolysaccharide with polyrhamnose 41,42. The expression of polyrhamnose on the surface of E. coli was confirmed by anti-GAC antibodies that recognize the GAC polyrhamnose backbone (Supplementary Fig. 11). Flow cytometric analysis revealed that AtlA-GFP only bound to polyrhamnose-producing E. coli and not to the parental strain (Fig. 4c). These data unambiguously demonstrate that the SCC polyrhamnose backbone is sufficient to confer AtlA binding. GFP alone did not bind to the recombinant bacteria (Fig. 4c).
GroP and the Glc side-chains hinder AtlA-GFP binding to the polyrhamnose backbone
To investigate how the presence of the SCC side-chain substituents affects recognition of the polyrhamnose backbone by AtlA, we compared AtlA binding affinities to various SCCs using fluorescence polarization anisotropy. The SCC variants prepared from WT, ΔsccH, ΔsccN, and ΔsccNΔsccP were conjugated with a fluorescent tag (ANDS). The colorless AtlA-GFP variant protein, AtlA-cGFP, was incubated with the fluorescently labeled SCCs at varying concentrations (Fig. 4d). We observed that the binding of the ΔsccNΔsccP SCC is the strongest, with an apparent Kd = 0.9 µM (95% confidence interval: 0.6-1.2), which is in the range of the binding affinity of known lectins for complex glycans 43,44. While binding of the WT, ΔsccH, and ΔsccN SCCs was unable to reach saturation due to solubility constraints, they all demonstrate clear evidence of binding to AtlA-cGFP (Fig. 4d). The ΔsccN SCC has the second-highest affinity, estimated to be at least 12-fold weaker than ΔsccNΔsccP, with the ΔsccH and WT SCCs being indistinguishable under accessible conditions. Both SCCs have at least a 25-fold lower affinity compared to SCC isolated from ΔsccNΔsccP bacteria. The differences in binding of the SCC variants suggest that the primary recognition site of the AtlA Bsp repeat domain is unmodified polyrhamnose, and the addition of branching structures decreases binding affinity, presumably due to steric hindrance. The differences in binding between the ΔsccNΔsccP and ΔsccN SCCs further reinforces our hypothesis that both SccN and SccP provide the Glc donor for modification of SCC with the side-chains.
To further explore differences in binding of AtlA-GFP to the SCCs extracted from WT, ΔsccH, and ΔsccN, we employed analytical ultracentrifugation. Continuous c(s) distribution analysis estimated AtlA-GFP to have a molecular weight of ∼110 kDa, confirming that the protein fusion remains mostly monomeric at the concentration range being studied (Fig. 4e). This verifies that assembly is polysaccharide-dependent and not a function of the GFP tag. Upon addition of the WT, ΔsccH, or ΔsccN SCCs, higher-order complexation was observed between 10-20 S (Fig. 4e). The width of the distribution and the high s values of the largest species imply that many different permutations are occurring simultaneously. There is likely an equilibrium of different combinations: one AtlA protein bound to multiple SCCs, multiple proteins bound to a single SCC, and higher-order configurations of SCCs and AtlA proteins bridging and/or daisy-chaining together to form very large assemblies. Because c(s) is a weight-averaged model, molecular weight estimates are not meaningful for these types of distributions. However, the experiments demonstrate that WT SCC produces, on average, a lower-sized complex than either the ΔsccH or ΔsccN polysaccharides.
To better understand overall complexation, the same data were analyzed using wide distribution analysis (Fig. 4f). Each sample resolved into a more Gaussian distribution, with a weight-averaged sedimentation coefficient of 5.5 S for AtlA-GFP alone, 10.6 S for the WT SCC, 12.3 S for the ΔsccH SCC, and 14.5 S for the ΔsccN SCC. Based on these values, wide distribution analysis also confirms the formation of smaller complexes in the presence of WT SCCs compared to the ΔsccH and ΔsccN polysaccharides. Thus, our results indicate that AtlA-GFP binds to the polyrhamnose backbone of SCC, and the modification of SCC with GroP and the Glc side-chains obstruct AtlA-GFP attachment to the polyrhamnose backbone.
SCC is highly heterogeneous with regard to GroP modification
AtlA localization studies together with the analysis of AtlA-GFP binding affinity are consistent with the idea that S. mutans equatorial sites and poles contain SCCs deficient in either GroP or Glc-GroP, whereas the sidewalls carry the fully mature SCC species decorated with Glc-GroP. Importantly, carbohydrate analysis of SCCs extracted from S. mutans WT, ΔsccH and ΔsccH:psccH cell walls by mild acid hydrolysis established that deletion of sccH has no significant effect on total SCC content (Supplementary Fig. 9b). This fact excludes the possibility that the mislocalized binding of AtlA along the entire surface of ΔsccH bacteria is due to increased expression of SCC within the cell wall. To further test our hypothesis that S. mutans peptidoglycan is decorated with the SCC variants lacking either GroP or Glc-GroP, we separated the extracted SCCs using anion-exchange chromatography. The majority of WT SCC is negatively charged and binds tightly to the anion-exchange column, eluting as a broad peak (Fig. 4g). However, a substantial portion (∼15-20 %) is neutral, with very low phosphate content, and does not bind to the anion-exchange column. Our previous work on the GAC revealed similar results 28, suggesting that streptococcal species produce different forms of the cell wall polysaccharides. As we previously reported, the phosphate content in GAC and SCC is an indication of the presence of the GroP modification in the polysaccharide 28. Analysis of the glycosyl composition in the two fractions revealed that the column-bound fraction contains more Glc relative to Rha than the unbound fraction (Fig. 4h). As expected, SCC extracted from ΔsccH elutes as a single peak of neutral SCC, lacking detectable phosphate (Supplementary figure 12).
To further characterize the heterogeneity of the SCC variants, we examined the electrophoretic mobility of ANDS-conjugated polysaccharides extracted from WT, ΔsccH, ΔsccN, ΔsccP and ΔsccNΔsccP (Fig. 4i). Since ANDS introduces a negative charge to SCC, this fluorescent tag allows examination of electrophoretic mobility of neutral polysaccharides extracted from the GroP-deficient mutants: ΔsccH, ΔsccN and ΔsccNΔsccP. We observed the distinctive “laddering” of bands in the SCCs extracted from WT and ΔsccP, indicating the high level of heterogeneity of the polysaccharides. As expected and in agreement with the anion-exchange chromatography analysis, the SCCs extracted from ΔsccH, ΔsccN and ΔsccNΔsccP migrated as a single band. Altogether, these data indicate that S. mutans WT produces SCC variants with different degrees of GroP modification.
Z-ring positioning is controlled by GroP modification
The morphological abnormalities of ΔsccH and ΔsccN suggest a role for GroP modification of SCC in regulating either the assembly or correct positioning of the Z-ring. To test this hypothesis, we expressed FtsZ fused with tagRFP (FtsZ-tagRFP) from its chromosomal locus as the only copy in WT and ΔsccH cells (the ftsZ-tagRFP and ΔsccH ftsZ-tagRFP strains). To identify the localization of the GroP-deficient SCCs on the surface of S. mutans, we added AtlA-GFP exogenously to the ftsZ-tagRFP and ΔsccH ftsZ-tagRFP strains. As expected, we observed both FtsZ-tagRFP and AtlA-GFP at the mid-cell in newborn WT cells (Fig. 5a, stage 1). In the cells beginning to elongate, the AtlA-GFP signal splits into two parallel bands that move away from the division site where a single FtsZ-ring is present (Fig. 5a, stages 2 and 3). During middle-to-late cell division stage, a part of FtsZ splits into two rings and migrates to the mid-cell regions of the newly forming daughter cells, resulting in a distinctive three-band pattern (Fig. 5a, stage 4). AtlA-GFP was already localized at these sites, suggesting that the wall regions carrying the GroP-deficient SCCs are present at the equators of daughter cells prior to the arrival of FtsZ (Fig. 5a, stage 4). This localization pattern is consistent with our idea that the equatorial rings contain immature SCCs that lack GroP. Interestingly, during the late cell division stage, the AtlA-GFP signal was detected at the constricting septum (Fig. 5a, stage 5), indicating that the GroP-deficient SCCs are inserted in the septal cell wall during the pole maturation.
In the ΔsccH ftsZ-tagRFP cells, the AtlA-GFP signal was evenly distributed along the cell surface, reflecting the presence of the GroP-deficient SCCs throughout the whole cell wall. While FtsZ was able to assemble into the Z-ring structures in the mutant cells, some Z-rings were often displaced from the cell center or not perpendicular to the cell’s axis (Fig. 5b and Supplementary figure 13a). These data indicate that the morphological defects of ΔsccH arise from a misplacement of the Z-ring.
GroP modification regulates MapZ-ring positioning
The above results suggest that the correct positioning of Z-ring at the mid-cell depends on the presence of the GroP-deficient SCCs in the equatorial ring. Intriguingly, the cell shape alterations, together with the Z-ring misplacement observed in ΔsccH, are reminiscent of S. pneumoniae and S. mutans mutants lacking MapZ 7-10, implicating the GroP-deficient SCCs in the recruitment of MapZ to the equatorial ring. To test this hypothesis, we expressed the fusion of MapZ with GFP (MapZ-GFP) in the WT and ΔsccH genetic backgrounds resulting in the mapZ-GFP and ΔsccH mapZ-GFP strains. To analyze the correlation between MapZ and FtsZ, we generated the MapZ-GFP/FtsZ-tagRFP double expressing strains both in the WT and ΔsccH genetic backgrounds resulting in the ftsZ-tagRFP mapZ-GFP and ΔsccH ftsZ-tagRFP mapZ strains. The localization of the GroP-deficient SCCs on the surface of the mapZ-GFP strain was monitored by labeling bacteria with AtlA-tagRFP protein (the tagRFP fusion with the N-terminal Bsp repeats domain of AtlA), which was added exogenously. As expected from previous studies 6-10, in the WT newborn cells, the MapZ-ring coincides with the FtsZ-ring at mid-cell (Fig. 5c, stage 1). As cell division progresses, MapZ migrates as a pair of rings, parallel to the respective equators of newly forming daughter cells, while a single FtsZ-ring remains at the mid-cell of the parental cell (Fig. 5c, stages 2 and 3). At a later cell division stage, the majority of the FtsZ arrives at the future division sites in the daughter cells, where it co-localizes with MapZ (Fig. 5c, stage 4). AtlA-tagRFP associated with the WT cells with patterns very similar to that observed with AtlA-GFP being enriched at the poles and mid-cell regions (Fig. 5d). Importantly, the MapZ signal co-localizes with GroP-deficient SCCs indicated by AtlA-tagRFP labeling at mid-cell regions in newborn and elongating WT cells (Fig. 5d), which is consistent with the idea that these immature SCCs are present in the equatorial rings.
Strikingly, in the ΔsccH (strains ΔsccH mapZ-GFP and ΔsccH ftsZ-tagRFP mapZ-GFP) cells, MapZ did not assemble into the characteristic ring-like structures, but instead the MapZ-GFP fluorescent signal was dispersed throughout the membrane of the mutants (Fig. 5e and Supplementary figure 13b). As expected from the proposed function of MapZ in guiding FtsZ positioning 7-10, Z-rings were mislocalized in the ΔsccH cells (Fig. 5e). These results indicate that the enrichment of the GroP-deficient SCCs at the equatorial rings drives the recruitment and assembly of MapZ into the ring-like structures. The underlying mechanism likely relies on an exclusion strategy whereby decoration of SCC with GroP at distinct cellular positions provides the molecular signal to exclude recruitment of MapZ and FtsZ to these sites.
Discussion
Despite years of intensive research, it remains unclear how oval-shaped Gram-positive bacteria generate equally sized daughter cells after division. This work uncovers a mechanism of molecular exclusion to position the cell division machinery in the human pathogen S. mutans. Our results are consistent with a model in which GroP-deficient SCCs are enriched at the equatorial rings and poles. Such controlled distribution of the polysaccharides provides the molecular cues for the simultaneous recruitment of cell division machinery as well as proper daughter cell separation.
Rha-containing cell wall polysaccharides are expressed by a wide variety of streptococcal, lactococcal and enterococcal species 29. We recently reported the discovery of GroP modification in the S. mutans and GAS cell wall polysaccharides, SCC and GAC, respectively 28. This modification is likely present in other streptococci since homologs of sccH, which encodes the S. mutans GroP transferase, is highly conserved among streptococcal species except for the Streptococcus mitis group 28. Structural studies of the cell wall polysaccharides isolated from streptococcal species, including S. mutans, proposed that each disaccharide repeating unit of the polyrhamnose backbone is modified with glycosyl side-chains 29. Previously, we have demonstrated that in the GAS, the GlcNAc side-chains of GAC are decorated with GroP 28. Here, we report that the side-chains of SCC are similarly decorated by GroP in S. mutans. Moreover, we now demonstrate that the structural architecture of Rha-containing cell wall polysaccharides has important functional implications for regulation of cell division in streptococci.
We observe that the GroP- and Glc side-chain-deficient mutants of S. mutans display severely impaired cell division, as represented by a large fraction of unequally sized cells. The morphological changes are accompanied by reduced viability and enhanced susceptibility to autolysis. These intriguing phenotypes prompted a search for responsible cell division proteins and autolysins. Cell wall pulldown studies, together with co-sedimentation analysis using SCC-depleted cell walls, identified SCC as the only detectable cell wall binding receptor for AtlA. This autolysin is involved in cell separation of daughter cells after cell division, autolysis, bacterial competence, and biofilm formation in S. mutans 31-34,45. Fluorescence anisotropy experiments conclusively demonstrate that AtlA-GFP binds to Glc- and GroP-modified SCCs, but the binding affinity for these variants is significantly lower than for those lacking the decorations. The analytical ultracentrifugation studies further revealed that all analyzed SCC variants, including the WT SCCs, interact with AtlA to form higher-order complexes. However, modification of SCC with either GroP or the Glc side-chains reduces the extent of the protein oligomerization. We show that the AtlA N-terminal domain which is composed of Bsp repeats, targets specifically the polyrhamnose backbone of SCC. This finding allowed us to employ a fusion of the AtlA N-terminal domain with a fluorescent protein as a tool to map the topological arrangements of different SCC species within the bacterial cell wall.
Using immunofluorescent microscopy we demonstrate that AtlA and AtlA-GFP are evenly distributed along the cell surface of the mutants lacking GroP and Glc side-chains, indicating that peptidoglycan is decorated with SCCs throughout the whole cell surface of S. mutans. Importantly, the absence of binding of AtlA-GFP to the ΔrgpG cells eliminates the possibility that AtlA recognizes additional structures on the bacterial surface. Finally, expression of the SCC polyrhamnose backbone is sufficient to confer AtlA binding, as demonstrated by expression in the non-natural host E. coli. The autolysin was found to be responsible for enhanced susceptibility to autolysis of the GroP- and Glc side-chain-deficient mutants. In newborn WT cells, AtlA is specifically targeted to cell poles, where the enzyme hydrolyzes peptidoglycan to allow daughter cell separation, and to cell equators. These observations lead to the conclusion that the mislocalization of AtlA and its strong binding to the cell wall causes the enhanced cell lysis of the mutants.
During the elongation phase, the SCC species labeled with the fluorescent AtlA fusion move from mid-cell of the parental cell, where the Z-ring is localized, toward the cell equators of the newly forming daughter cells. Subsequently, these SCCs arrive at the new division site before FtsZ. The apparent movement of the AtlA-labeled SCCs is the result of the newly synthesized cell wall being added at the septum, pushing the zone with these specific SCCs away from the cell division site. Furthermore, the AtlA-labeled SCCs co-localize with MapZ during the division cycle, indicating the presence of the distinct form of SCC in the equatorial rings. Characterization of SCCs extracted from S. mutans cell wall revealed a high degree of heterogeneity with a significant portion of SCC containing low or no GroP. These results, combined with the observed difference in AtlA localization in WT versus GroP-deficient bacteria, are consistent with the idea that peptidoglycan in the equatorial rings and cell poles is decorated with newly synthesized SCCs lacking GroP modifications. Note, the results do not exclude the possibility that these SCCs are also deficient for the Glc side-chain.
The highly organized spatial distribution of the GroP-deficient SCC variants further implies that the synthesis of the equatorial rings and cell poles requires a specific modulation of the divisome and/or elongasome complexes to produce cell wall decorated with immature SCCs at the early and final stages of cell division. In line with this idea, our co-localization analysis revealed that the translocation of FtsZ from the parental septum to the equators of the daughter cells coincides with the re-appearance of the AtlA-GFP-labeled SCCs at the parental septum, suggesting that the GroP-deficient cell wall material is incorporated into the septal wall at the last step of septum closure. This observation is in agreement with the time-lapse ultrastructural reconstruction of dividing E. faecalis indicating that the synthesis of septum occurs in two separate events — early in the division and after the elongation phase 1.
In ovococci, the peptidoglycan architecture at the equatorial rings has been proposed to direct the positioning of the cell division machinery through MapZ 7,8. Our analysis of MapZ localization in the GroP-deficient mutant revealed its impaired alignment with the equatorial rings. Interestingly, a similar localization defect was observed for MapZ lacking its extracellular domain in S. pneumoniae 7. Furthermore, we found that the delocalization of MapZ in the mutant cells causes a mis-orientation of the Z-rings, indicating that the cell shape deformations observed in the GroP-deficient mutants arise from a defect in septum placement. These findings highlight the pivotal role of GroP modification of SCC in cell division, leading to a model wherein immature SCCs, serve as landmarks for the recruitment of MapZ to equatorial rings and AtlA to equatorial rings and cell poles (Fig. 5f). Thus, this simple mechanism unifies septum positioning with its subsequent splitting. The importance of AtlA binding to equatorial rings is unclear at the moment. Early in the cell division cycle, AtlA might participate in the shaping of the new equatorial rings of the daughter cells by splitting the parental equatorial ring (Fig. 5f). A previous report, using electron microscopy imaging, has identified the equatorial rings as the zones of autolysin activity in E. faecalis 46. However, since the deletion of AtlA does not affect S. mutans cell shape, other mechanisms besides AtlA might be responsible for the generation of the daughter equatorial rings.
How MapZ recognizes cell wall decorated with immature SCCs is currently unknown but under investigation. Since MapZ homologs are also present in the species of the S. mitis group that do not express Rha-containing cell wall polysaccharides 7,8, it suggests that, in contrast to AtlA, MapZ does not interact directly with the cell wall polysaccharides. Note, that the morphological phenotype of GroP-deficient S. mutans does not depend on the specific glycosyl side chain to which GroP is attached since the mutant phenotype is complemented by the plasmid-based expression of the GAC-specific side-chain. This observation suggests that a major function of GroP in streptococcal cell wall is to provide a negative charge. Phosphate groups in WTA have been proposed to affect the packing density and rigidity of the cell wall due to electrostatic repulsion between the polysaccharides 47. Thus, it is possible that a cue for recognition of the equatorial ring by MapZ is an alteration in peptidoglycan density.
The Rha-containing cell wall polysaccharides are the functional homologs of canonical WTAs in streptococci because, similar to WTAs, they play significant roles in metal ion homeostasis, antimicrobial resistance 28, cell division and autolysis. The exact molecular mechanism by which WTAs regulate cell division and autolysis is not known. Much like GroP modification, WTAs are required for proper localization of autolysins by restricting the binding of autolysins from the entire bacterial surface and directing them to the specific cell wall sites where they are enzymatically active 13,15,17. Considering the similar function of WTAs and GroP modification of SCC, it is plausible that the here-described mechanism of the positioning of cell division proteins is widespread in Gram-positive bacteria. However, cell separation autolysins and the proteins bridging the Z-ring to cell wall likely differ between species.
Methods
Bacterial strains, growth conditions and media
All plasmids, strains and primers used in this study are listed in Supplementary Tables 1 and 2. Streptococci and E. faecalis were grown in BD Bacto Todd-Hewitt broth supplemented with 1% yeast extract (THY) without aeration at 37°C. S. mutans strains were grown either on THY or brain heart infusion (BHI) agar at 37°C with 5% CO2. E. coli strains were grown in Lysogeny Broth (LB) medium or on LB agar plates at 37°C. When required, antibiotics were included at the following concentrations: ampicillin at 100 μg mL-1 for E. coli; erythromycin at 5 μg mL-1 for S. mutans; chloramphenicol at 10 µg mL-1 for E. coli and 2 µg mL-1 for S. mutans; spectinomycin at 500 μg mL-1 for S. mutans; kanamycin at 300 μg mL-1 for S. mutans.
Construction of mutant strains
To delete sccN, S. mutans Xc chromosomal DNA was used as a template for the amplification of two DNA fragments using two primers pairs: sccNup-BglII-f/sccNup-SalI-r and sccNdown-BamHI-f/sccNdown-XhoI-r (Supplementary Table 2). The first PCR product was digested with BglII/SalI and ligated into BglII/SalI-digested pUC19BXspec 30. The resultant plasmid was digested with BamHI/XhoI and used for ligation with the second PCR product that was digested with BamHI/XhoI. The resultant plasmid, pUC19BXspec-sccN, was digested with BglII and XhoI to obtain a DNA fragment containing the nonpolar spectinomycin resistance cassette flanked with the sccN upstream and downstream regions. The DNA fragment was transformed into S. mutans Xc by electroporation. The mutants were isolated as described below. The plasmids for the deletion of sccP and atlA were constructed using the same strategy described for sccN deletion with primer pairs listed in Supplementary Table 2.
To delete rgpG and smaA in the WT strain, smaA in the ΔatlA and ΔsccN backgrounds, sccP in the ΔsccN background, and atlA in the ΔsccN and ΔsccH backgrounds, we used a PCR overlapping mutagenesis approach, as previously described 28. Briefly, 600-700 bp fragments both upstream and downstream of the gene of interest were amplified with designed primers that contained 16-20 bp extensions complementary to the nonpolar antibiotic resistance cassette (Supplementary Table 2). The nonpolar spectinomycin, kanamycin, and erythromycin resistance cassettes were PCR-amplified from pLR16T, pOSKAR, and pHY304 (Supplementary Table 1), respectively. The two fragments of the gene of interest and the fragment with the antibiotic resistance cassette were purified using the QIAquick PCR purification kit (Qiagen) and fused by Gibson Assembly (SGA-DNA). A PCR was then performed on the Gibson Assembly sample using primers listed in Supplementary Table 2 to amplify the fused fragments.
To construct S. mutans strains expressing FtsZ fused at its C-terminus with a monomeric red fluorescent protein (tagRFP) 48, we replaced ftsZ with ftsZ-tagRFP at its native chromosome locus. A nonpolar kanamycin resistance cassette was inserted downstream of ftsZ-tagRFP to allow the selection of recombinant bacteria. The fragment encoding tagRFP was PCR-amplified from pTagRFP-N (Evrogen). The fragments of ftsZ, tagRFP, kanamycin resistance cassette, and the ftsZ downstream region were PCR-amplified using primers listed in Supplementary Table 2, purified and assembled as described above. S. mutans strains expressing MapZ fused at its N-terminus with a superfolder green fluorescent protein (GFP) 36 were constructed similarly, except that a nonpolar spectinomycin resistance cassette followed by the GAS mapZ promoter was inserted upstream of the mapZ fusion to allow the mapZ expression and selection of recombinant bacteria. The fragments of the mapZ upstream region, spectinomycin resistance cassette, GFP, and mapZ were PCR-amplified using primers listed in Supplementary Table 2. The assembled PCR fragment was transformed into corresponding S. mutans strains by electroporation. The transformants were selected either on THY or BHI agar containing the corresponding antibiotic. Double-crossover recombination was confirmed by PCR and Sanger sequencing using the primers listed in Supplementary Table 2.
Complementation of ΔsccN
To complement ΔsccN, sccN was amplified from S. mutans Xc chromosomal DNA using primer pair sccN-HindIII-f/sccN-BglII-r, digested using HindIII/BglII, and ligated into HindIII/BglII-digested pDC123, yielding psccN. To replace the SCC side-chain with the GAC side-chain, a part of the GAC operon required for the addition of the GlcNAc side-chains and GroP 30 was expressed on pDC123 in ΔsccN. GAS 5448 genomic DNA was used to amplify the gacHIJKL region using primer pairs A109-f/A101-r (Supplementary Table 2). The PCR fragment was digested using XhoI/BamHI, and ligated into XhoI/BamHI-digested pDC123, yielding pgacHIJKL. All plasmids were confirmed by sequencing analysis (Eurofins MWG Operon) before electroporation into S. mutans. We found that one E. coli clone carrying pgacHIJKL acquired a frameshift resulting in a stop codon at leucine 49 in GacI. The plasmid was designated pgacHI*JKL and used as a negative control in our experiments. Plasmids were transformed into ΔsccN by electroporation. Chloramphenicol resistant single colonies were picked and checked for the presence of psccN or pgacHIJKL by PCR, yielding strains ΔsccN:psccN, ΔsccN:pgacHIJKL and ΔsccN:pgacHI*JKL.
Construction of the plasmids for expression of AtlA-GFP, AtlA-cGFP, AtlA-tagRFP and GFP
To create a vector for expression of the AtlA N-terminal Bsp domains fused with a superfolder GFP (AtlA-GFP fusion protein), two overlapping amplicons were generated. The first fragment encoding the N-terminal Bsp domains was amplified from S. mutans Xc chromosomal DNA using the primer pair atlA_fus-F and atlA_fus-R. The second fragment encoding GFP was amplified from pHR-scFv-GCN4-sfGFP-GB1-NLS-dWPRE (Supplementary Table 1) using the primer pair gfp_fus-F and gfp_fus-R. The PCR products were ligated into NcoI/HindIII-digested pRSF-NT vector (Supplementary Table 1) using Gibson assembly, resulting in pKV1527 for expression of AtlA-GFP.
To create a vector for expression of the AtlA N-terminal Bsp domains fused with colorless GFP (AtlA-cGFP fusion protein), two PCR products were amplified using primer pairs atlA_fus-F/Y66L_R and Y66L_F/gfp_fus-R, and pKV1527 as a template to introduce Y66L mutation into GFP 49.
To create a vector for expression of the AtlA N-terminal Bsp domains fused with tagRFP (AtlA-tagRFP fusion protein), two PCR products were amplified using primer pairs atlA_fus-F/atl_tRFP_R and tRFP_fusF/tRFP_fusR using S. mutans Xc chromosomal DNA and pTagRFP-N, respectively, as the templates. The corresponding PCR products were ligated into NcoI/HindIII-digested pRSF-NT vector using Gibson assembly resulting in pKV1556 and pKV1572.
To create a vector for expression of GFP, a PCR product was amplified using primer pair sfGFP_BspH/gfp_fus-R and pKV1527 as a template. The PCR product was digested with BspHI/HindIII and ligated into the NcoI/HindIII-digested pRSF-NT vector resulting in pKV1532.
Protein expression and purification
To purify AtlA-GFP, AtlA-cGFP, AtlA-tagRFP and GFP, E. coli Rosetta (DE3) cells carrying the respective plasmids were grown in LB at 37°C to OD600 = 0.4-0.6 and induced with 0.25 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) at 27°C for approximately 4 hours. Bacteria were lysed in 20 mM Tris-HCl pH 7.5, 300 mM NaCl by a microfluidizer cell disrupter. The proteins were purified by Ni-NTA chromatography followed by size exclusion chromatography (SEC) on a Superdex 200 column in 20 mM HEPES pH 7.5, 100 mM NaCl.
Viability assay
Exponentially growing bacteria (OD600 of 0.5) were pushed ten times through a 26G 3/8 syringe to break bacterial clumps. Bacteria were serially diluted in phosphate-buffered saline (PBS) and plated on THY agar for enumeration.
Triton X-100-induced autolysis assay
Overnight cultures of S. mutans were diluted (1:100) into fresh THY broth and grown to an OD600 of 0.5. The autolysis assay was primarily performed, as outlined in 50. Cells were allowed to autolyze in PBS containing 0.2% Triton X-100. The autolysis was monitored after 2, 4, and 21 hours as a decrease in OD600. Results were normalized to the OD600 at time zero (OD600 of 0.5).
Isolation of cell walls and sacculi
The cell walls were isolated from exponential phase cultures by the SDS-boiling procedure and lyophilized, as previously described 30. The sacculi were obtained using the same protocol except that the bead beating step was omitted. The cell wall and sacculi were free of lipoteichoic acid (LTA), proteins, lipids, and nucleic acids.
Binding of AtlA-GFP to intact bacteria and cell wall material
To study AtlA-GFP binding to intact bacteria, 10 mL of the overnight-grown bacteria were washed twice with PBS, resuspended in 1 mL of PBS, and incubated with 0.1 mg mL-1 AtlA-GFP for 1 hour with agitation. As a control, GFP of the same concentration was used in parallel with each experiment. Sample aliquots were assayed to determine the total fluorescence. Then samples were centrifuged (16,000 g, 3 min), and 100 µL of the supernatant was assayed for fluorescence. To determine the fluorescence of the pellet, the supernatant fluorescence was subtracted from the total fluorescence of the sample. Controls without bacterial cells indicated that the sedimentation of AtlA-GFP under these conditions was negligible. Data are presented as a percentage of fluorescence of the pellet normalized to the total fluorescence of the sample.
To examine AtlA-GFP binding to purified cell walls, 0.5 mg of lyophilized cell wall was incubated with 0.1 mg mL-1 AtlA-GFP in 0.5 mL of PBS. The experiment was conducted as described above.
Pulldown of cell wall-associated proteins
S. mutans (1 L) were grown to an OD600 of 1.0, collected by centrifugation (5,000 g, 10 min), washed three times with PBS, and resuspended in 25 mL of urea solution (8 M urea, 20 mM Tris-HCl pH 7.5, 150 mM NaCl). The sample was rotated at room temperature for 1 hour, and then centrifuged (3,200 g, 10 min). The supernatant was dialyzed overnight at 4°C to remove urea and centrifuged again (3,200 g, 10 min). The supernatant was transferred to a fresh centrifuge tube and incubated with 10 mg of lyophilized cell wall with rotation for two hours. The cell wall was collected by centrifugation (3,200 g, 10 min) and washed three times with PBS. The proteins, retained in the cell wall, were dissolved in 0.5 mL of SDS sample buffer, and separated on 10% SDS-PAGE. Protein identification was performed at the Proteomics Core Facility (University of Kentucky) by liquid chromatography with tandem mass spectrometry (LC-MS/MS) analysis using an LTQ-Orbitrap mass spectrometer (Thermo Fisher Scientific) coupled with an Eksigent Nanoflex cHiPLC system (Eksigent) through a nanoelectrospray ionization source. The LC-MS/MS data were subjected to database searches for protein identification using Proteome Discoverer software V. 1.3 (Thermo Fisher Scientific) with a local MASCOT search engine.
Release of SCCs from cell wall by mild acid hydrolysis
SCC was released from purified cell walls by mild acid hydrolysis following N-acetylation, as previously described for WTA of Lactobacillus plantarum 51 with some modifications. N-acetylation was conducted with lyophilized cell wall (4 mg) and 2 % acetic anhydride in 1 mL of saturated NaHCO3. After incubation at room temperature overnight, the reactions were diluted with 2 vol water, and sedimented at 50,000 g, 10 min. The cell walls were further washed by resuspension with 3 mL water followed by sedimentation at 50,000 g, 10 min, three times. N-acetylated cell walls were resuspended with 0.02 N HCl (0.2 mL), and heated to 100°C for 20 min. The reactions were cooled on ice, neutralized by the addition of 4 µL 1 N NaOH, and sedimented at 50,000 g, 10 min. The supernatant fraction was removed and reserved. The pellet was resuspended in 0.2 mL water and re-sedimented. The supernatant fractions were combined and either analyzed or purified further by a combination of SEC and ion-exchange chromatography.
Fractionation of SCCs on DEAE-Toyopearl
SCCs were released from purified cells walls by mild acid hydrolysis and fractionated on BioGel P150 equilibrated in 0.2 N sodium acetate, pH 3.7, 0.15 M NaCl, as previously described using BioGel P10 28. The fractions containing SCCs were combined and concentrated by centrifugation over an Amicon Ultra - 15 Centrifugal Filter (KUltracel -3K). The retentate was desalted by repeated cycles of dilution with water and centrifugation. The concentrated SCCs (∼0.5 mL) were applied to a 1×18 cm column of TOYOPEARL DEAE-650M (TOSOH Bioscience), equilibrated in 10 mM Tris-Cl, pH 7.4, and fractions of 2 mL were collected. After 20 fractions, the column was eluted with an 80 mL gradient of NaCl (0-0.5 M). Appropriate aliquots were analyzed for Rha and Glc by anthrone assay.
Derivatization of SCCs with 7-amino-1,3-naphthalenedisulfonic acid (ANDS)
SCCs purified by a combination of SEC and ion-exchange chromatography were fluorescently tagged at the reducing end by reductive amination with ANDS as previously described 52. Reaction mixtures contained 30-100 nmol of SCCs (lyophilized), 0.75 mM ANDS and 0.5 M NaCNBH4 in 0.05 mL acetic acid/DMSO at 7.5/50 (%). Following overnight incubation at 37°C, the reactions were desalted by centrifugation on an Amicon Ultra Centrifugal Filter (3,000 NMWL). Derivatized SCCs were further purified by SEC over a Superdex 75 10/300 GL column (GE Healthcare Bio-Sciences AB).
Glycerol and phosphate assays
The total phosphate content of SCCs was determined by the malachite green method following digestion with perchloric acid, as previously described 28. To determine glycerol, SCCs were incubated with 2 N HCl at 100°C for 2 hours. The samples were neutralized with NaOH in the presence of 62.5 mM HEPES pH 7.5. Glycerol concentration was measured using the Glycerol Colorimetric assay kit (Cayman Chemical) according to the manufacturer’s protocol.
Modified anthrone assay
Total Rha and Glc contents were estimated using a minor modification of the anthrone procedure. Reactions contained 0.08 mL of aqueous sample and water and 0.32 mL anthrone reagent (0.2 % anthrone in concentrated H2SO4). The samples were heated to 100°C, 10 min, cooled in water (room temperature), and the absorbance at 580 nm and 700 nm was recorded. The chromophore produced from Rha has a relatively discreet absorbance maximum and is essentially zero at 700 nm. The absorbance of the Glc chromophore at 700 nm is approximately equal to its absorbance at 580 nm. Therefore the contribution of Glc to the absorbance at 580 nm can be estimated from its absorbance at 700 nm and subtracted from the absorbance at 580 nm to obtain the Rha-specific signal. Rha and Glc concentrations were estimated using L-Rha and D-Glc standard curves, respectively.
Glycosyl composition analysis
Glycosyl composition analysis of SCC samples was performed at the Complex Carbohydrate Research Center (University of Georgia, Athens, GA) by combined gas chromatography/mass spectrometry (GC-MS) of the per-O-trimethylsilyl derivatives of O-methyl glycosides of the monosaccharides produced by acidic methanolysis as described previously 30. Likewise, a similar assay was performed following in-house pretreatment with HF, as described below. Appropriate aliquots were supplemented with 2 nmol inositol (internal standard), taken to dryness, and incubated with 25 µL 51% HF, 4°C, 16 hours. Following HF treatment, the samples were evaporated under a stream of air (the evaporating HF was captured by bubbling through a trap containing 1 M KOH), dissolved in water, transferred to screw-cap tubes equipped with Teflon lined caps and dried again. 0.2 mL 1 N HCl in methanol (formed by the drop-wise addition of acetyl chloride into rapidly-stirring anhydrous methanol) was added, the tubes were tightly sealed and incubated at 80°C, 16 hours. Following methanolysis, the reactions were neutralized with ∼3-5 mg AgCO3, centrifuged, and the supernatant transferred to a fresh tube. To re-N-acetylate amino sugars, the samples were taken to dryness under a stream of air, evaporated out of 0.5 mL of methanol, and re-dissolved in 0.1 mL of methanol containing 10 % pyridine and 10 % acetic anhydride. The reactions were dried, trimethylsilylated with 25 μL Tri-Sil TH (Sigma Aldrich), and analyzed by methane chemical ionization GC-MS using a Thermo ISQ mass spectrometer interfaced with a gas chromatograph, equipped with a 15 m Equity 1701 glass capillary column and helium carrier gas.
GlcNAc analysis
GlcNAc content was assayed using the Megazyme Glucosamine Assay Kit according to the manufacturer’s instructions with some minor modifications. Partially purified polysaccharide (∼40-60 nmol Rha) was hydrolyzed in 40 µL 2 N HCl, 100°C, 2 hours, neutralized with 10 N NaOH (to ∼pH 7.0 by pH paper) and adjusted to a final volume of 50 µL with water. The acid hydrolysis de-acetylates GlcNAc to generate glucosamine. An aliquot (5 µL) of the neutralized hydrolysate was diluted with 171 µL water and mixed with a total of 24 µL of Megazyme Glucosamine Assay Kit Enzyme Mix. Absorbance at 340 nm was recorded after incubation at room temperature for 40 min. Glucosamine content was determined by comparison with a glucosamine standard curve and verified by comparison with GlcNAc standards treated similarly as the polysaccharide.
FACS analysis
AtlA-GFP (5 µL, 3.8 mg mL-1) or GFP (5 µL, 3.5 mg mL-1) were added to 100 µL of E. coli CS2775 41 or PHD136 [E. coli CS2775 harboring pRGP1 plasmid 41] (OD600 = 0.4). After 20 minutes of incubation on ice, the cells were centrifuged at 20,800 g, 5 min. The pellet was washed twice with PBS, resuspended in PBS, fixed with paraformaldehyde (4% final concentration), 4°C, 20 min, and then washed once with PBS. The cells were resuspended in 0.3% BSA in PBS and immediately analyzed by flow cytometry (BD LSRFortessa). Anti-GAC antibodies conjugated with FITC (ABIN238144, antibodies-online, titer 1:50) were used as a positive control to confirm polyrhamnose expression in E. coli strain PHD136. The FACS data were analyzed using flowJo version 10.
Scanning electron microscopy (SEM)
Exponentially growing bacteria (OD600 of 0.7) were harvested by centrifugation (3,200 g, 10 min), washed once with 20 mM HEPES, 0.5% BSA buffer (pH 8.0), resuspended in PBS, fixed with paraformaldehyde (4% final concentration), 4°C, 15 min, and then pipetted onto microscope slide cover glasses coated with poly-L-lysine. Following one hour incubation, the cover glasses were washed 3 times with PBS. Bacteria were dehydrated stepwise in a gradient series of ethanol (35%, 50%, 70%, and 96% for 20 min each and then 100% overnight), followed by critical point drying with liquid CO2 in a Leica EM CPD300. Samples were coated with about 5 nm of platinum controlled by a film-thickness monitor. SEM images were performed in the immersion mode of an FEI Helios Nanolab 660 dual beam system.
Fluorescent and differential interference contrast (DIC) microscopy
To determine the bacterial regions targeted by AtlA-GFP and AtlA-tagRFP, we utilized fluorescent microscopy. Exponentially growing bacteria (OD600 of 0.7) were fixed with paraformaldehyde (4% final concentration), 4°C, 15 min, pipetted onto microscope slide cover glasses coated with poly-L-lysine, and allowed to settle for one hour at room temperature. Bacteria were incubated with AtlA-GFP or AtlA-tagRFP (20 µg mL-1) for 15 min at room temperature. As a control, GFP of the same concentration was used in parallel with each experiment. The samples were washed four times with PBS, dried at room temperature, and mounted on a microscope slide with ProLong Diamond Antifade Kit with DAPI (Invitrogen). Samples were imaged on a Zeiss LSM 880 using Airyscan mode and a Leica SP8 equipped with 100X, 1.44 N.A. objective, DIC optics, and “lightning” post-processing. Images were processed with Airyscan processing and the “lightning” processing tool, respectively. Samples with S. mutans cells expressing MapZ-GFP and FtsZ-tagRFP were prepared similarly. They were imaged on a Leica SP8. Images were deconvolved using Huygens Professional software.
Immunofluorescent microscopy was used to monitor AtlA on the cell surface of S. mutans. Exponentially growing bacteria (OD600 of 0.8) were fixed with 2.5% (v/v) paraformaldehyde, 0.03% glutaraldehyde in 30 mM phosphate buffer (pH 8.0), transferred onto poly-L-lysine coated cover glasses and incubated at room temperature for one hour. The samples were washed three times with PBS and incubated in buffer containing 20 mM Tris-HCl pH 7.5, 10 mM EDTA, 50 mM Glc, and lysozyme (0.1 mg mL-1) for 30 min. After washing twice with PBS, the samples were air-dried, and dipped in cold methanol, -20°C, 5 min. The samples were blocked with 2% (w/v) bovine serum albumin in PBS (BSA-PBS), room temperature, 2 hours, and incubated with polyclonal anti-AtlA antibodies 33 (1:600) in BSA-PBS, 4°C, overnight. The samples were then washed 15 min five times with PBS+0.1% Tween-20 (PBST) buffer and incubated with Alexa Fluor 488-conjugated goat anti-rabbit antibodies (1:100) in BSA-PBS, room temperature, 2 hours. After extensive washing with PBST buffer, the specimens were air-dried and assembled on microscope slides mounted with ProLong Diamond Antifade Kit with DAPI. Micrographs were taken on a Leica SP8 confocal microscope.
To determine the length and width of cells, exponentially growing bacteria (OD600 of 0.7) were imaged on a Leica SP8 confocal microscope. ImageJ software was used to measure the sizes of cells.
Fluorescence anisotropy
Reactions (150 µL) containing 0.5 µM ANDS-labeled polysaccharide and 0-10 µM AtlA-cGFP were incubated at room temperature for 30 minutes in 20 mM Tris 7.5, 300 mM NaCl. The anisotropy was then measured on a Fluoromax-4 (Horiba) photon-counting steady-state fluorometer at 25°C using an excitation wavelength of 310 nm and the following emission at 450 nm with slit widths of 5 nm and an integration time of 1 second. All measurements are an average of three independent replicates. Curves terminate at 10 µM of AtlA-cGFP due to aggregation issues at higher protein concentration. The single-site total binding equation in Graphpad Prism 8 was used to fit the binding of the polysaccharide isolated from the ΔsccNΔsccP mutant. Lower-bound estimates of the Kd for the other three SCC species were determined by fitting to the binding equation after fixing the maximal binding limit, Bmax, to that of ΔsccNΔsccP.
Analytical ultracentrifugation
Sedimentation velocity (SV) experiments were performed in a Beckman ProteomeLab XL-I at 20°C using absorbance optics at 278 nm in an An-60Ti rotor at 32,000 rpm until complete sedimentation of sample occurred. The analysis was conducted using Sedfit 16.1c 53,54 using the c(s) data model and expressed using sedimentation coefficient distributions. Each fit rmsd was 0.005 or lower. GUSSI 1.4.1 55 was used for data visualization. SV data were also fitted using Wide Distribution Analysis (WDA) in SedAnal v7.11 56. WDA distributions were computed from 6.10 cm to 7.00 cm with an increment of 0.01 cm. The radial range plotted was 6.40 - 6.60 cm in 0.01 cm increments, with a 2% smoothing algorithm applied (equivalent to 4 scans). The weight-average sedimentation coefficient (sw) was computed using a range of 2-100 S. Partial specific volume (v-bar) was 0.725 ml g-1, and the solution density (ρ) was 0.998 g mL-1.
Statistical analysis
Unless otherwise indicated, statistical analysis was carried out on pooled data from at least three independent biological repeats. Statistical analysis of data was performed using one-way ANOVA, 2-way ANOVA, and two-tailed Student’s t-test as described for individual experiments. A P-value equal to or less than 0.05 was considered statistically significant.
Data availability
All data generated during this study are included in the article and supplementary information files or will be available from the corresponding author upon reasonable request.
Author contributions
SZ, CTC, JSR, SAC, AEY, ABH, NMvS, HCD, GIF, KVK, and NK designed the experiments. SZ, CTC, JSR, SAC, AEY, HCD, KVK, and NK performed functional and biochemical experiments. SZ and GIF performed microscopy analysis. NK, KVK, and NMvS constructed plasmids and isolated mutants. SZ, CTC, JSR, SAC, AEY, ABH, HCD, KVK, and NK analyzed the data. NK wrote the manuscript with contributions from all authors. All authors reviewed the results and approved the final version of the manuscript.
Competing interests
The authors declare no competing interests.
Acknowledgments
The authors thank Dr. Sang-Joon Ahn (University of Florida) for the kind gift of anti-AtlA antibodies, Dr. John F. Timoney (University of Kentucky) and Dr. Johannes Huebner (von Hauner Children’s Hospital, LMU) for providing S. equi and E. faecalis, respectively, Dr. Jeffrey M. Bosken and Dr. Edward D. Hall (University of Kentucky) for the use of the Thermo Scientific GC-MS instrument and Dr. Catalina Velez-Ortega (University of Kentucky) for the access to Leica SP8 confocal microscope. This work was supported by NIH grants R56 AI135021 from the NIAID and R01 DE028916 from the NIDCR (to NK), R01 GM094363 from the NIGMS (to ABH) and R01 DC014658 from the NIDCD (to GIF), Tenovus Scotland Large Research Grant T17/17 and University of Dundee Wellcome Fund 105606/Z/14/Z (to SAC and HCD), Wellcome and Royal Society Grant 109357/Z/15/Z (to HCD). Scanning electron microscopy was performed at the Electron Microscopy Center, which belongs to the National Science Foundation NNCI Kentucky Multiscale Manufacturing and Nano Integration Node, supported by ECCS-1542174. Carbohydrate composition analysis at the Complex Carbohydrate Research Center was supported by the Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Sciences, U.S. Department of Energy grant (DE-FG02-93ER20097) to Parastoo Azadi. The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.