An accessory protein required for anchoring and assembly of amyloid fibres in B. subtilis biofilmsmmi_

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1 Molecular Microbiology (2011) 80(5), doi: /j x First published online 4 May 2011 An accessory protein required for anchoring and assembly of amyloid fibres in B. subtilis biofilmsmmi_ Diego Romero, 1 Hera Vlamakis, 1 Richard Losick 2 and Roberto Kolter 1 * 1 Department of Microbiology and Molecular Genetics, Harvard Medical School, Boston, MA 02115, USA. 2 Department of Molecular and Cellular Biology, Harvard University, Cambridge, MA 02138, USA. Summary Cells within Bacillus subtilis biofilms are held in place by an extracellular matrix that contains cell-anchored amyloid fibres, composed of the amyloidogenic protein TasA. As biofilms age they disassemble because the cells release the amyloid fibres. This release appears to be the consequence of incorporation of D-tyrosine, D-leucine, D-tryptophan and D-methionine into the cell wall. Here, we characterize the in vivo roles of an accessory protein TapA (TasA anchoring/assembly protein; previously YqxM) that serves both to anchor the fibres to the cell wall and to assemble TasA into fibres. TapA is found in discrete foci in the cell envelope and these foci disappear when cells are treated with a mixture of D-amino acids. Purified cell wall sacculi retain a functional form of this anchoring protein such that purified fibres can be anchored to the sacculi in vitro. In addition, we show that TapA is essential for the proper assembly of the fibres. Its absence results in a dramatic reduction in TasA levels and what little TasA is left produces only thin fibres that are not anchored to the cell. Introduction When presented with a surface or an interface, bacteria will often grow as biofilms in which cells are held together by an extracellular matrix. For Bacillus subtilis this matrix is primarily composed of an exopolysaccharide and the amyloid protein TasA (Branda et al., 2001; Branda et al., 2006; Romero et al., 2010). Amyloid proteins are characterized by their ability to acquire a secondary structure enriched in b-sheets that facilitates the formation of Accepted 28 March, *For correspondence. rkolter@ hms.harvard.edu; Tel. (+1) ; Fax (+1) extremely stable fibres (Sunde et al., 1997; Kayed et al., 2003). While originally studied in the context of human pathologies, amyloid fibres are now recognized as ubiquitous extracellular structures in the microbial world, where they have functional roles (Fowler et al., 2007; Maury, 2009). These roles include mediating interactions with biotic and abiotic surfaces, raising aerial structures and providing structural integrity to the biofilm matrix (Collinson et al., 1993; Claessen et al., 2002; Gebbink et al., 2005; Barnhart and Chapman, 2006; Oh et al., 2007; Badtke et al., 2009). Because of their importance to bacterial physiology and their possible relevance as model systems for the study of human disease, the study of bacterial amyloid fibres has become an active area of research (Greenwald and Riek, 2010). While there is still much to understand about how amyloid fibres form and how they attach to the cell surface, some progress has been made by studying amyloid proteins produced by model bacteria. Polymerization of monomeric soluble subunits into complex and insoluble amyloid fibres can be aided by dedicated accessory proteins (Ferrone, 1999; Kayed et al., 2003; Epstein and Chapman, 2008; Tompa, 2009). This is the case for the curli fibres of Escherichia coli where the major structural subunit CsgA needs the accessory protein CsgB to polymerize in vivo (Hammar et al., 1996; Chapman et al., 2002; Barnhart and Chapman, 2006; Hammer et al., 2007). Streptomyces coelicolor also produces amyloid fibres. In this bacterium the fibres are composed of eight different chaplins, three of which can be covalently attached to the cell wall through the action of a sortase (Claessen et al., 2003; Claessen et al., 2006). Recently, it was found that some of the chaplins interact with cellulose in order to attach to the cell wall (de Jong et al., 2009). To date no nucleating factor for the formation of S. coelicolor fibres has been identified but proteins known as rodlins appear to facilitate the assembly process (de Jong et al., 2009). The TasA amyloid fibres provide structural integrity to B. subtilis biofilms (Romero et al., 2010). The fibres are anchored to cells and form a robust protein scaffold that holds the cells together. However, as biofilms senesce they gradually fall apart because the fibres are released from the cell. Recent results revealed some aspects of the mechanism of this disassembly. At late stages in a biofilm 2011 Blackwell Publishing Ltd

2 1156 D. Romero, H. Vlamakis, R. Losick and R. Kolter Fig. 1. tasa and tapa mutants do not complement each other extracellularly. A. Pellicle formation of wild-type and mutant strains. B. Pellicles from two strains mixed at a 1:1 ratio, designated with a + between the two genotypes. Pellicles were grown in MSgg medium for 24 h prior to imaging. life cycle, cells begin producing a mixture of D-amino acids that includes D-Tyr, D-Met, D-Leu and D-Trp. As these amino acids are incorporated into the peptidoglycan the TasA amyloid fibres are released from the cells, leading to biofilm disassembly (Kolodkin-Gal et al., 2010). Thus, it appears that the amyloid fibres are anchored to the cell wall and that this anchoring can be disrupted by incorporating unusual D-amino acids. Importantly, we obtained mutants that were resistant to the disassembling effect of D-Tyr (Kolodkin-Gal et al., 2010). These mutants carried altered forms of a protein, YqxM, that we had previously shown to be important for placing TasA in the extracellular matrix (Branda et al., 2006). TasA is encoded by tasa, the third gene in a three-gene operon. The two other proteins encoded in this operon are also involved in the process of amyloid fibre formation. The second gene encodes SipW, a signal sequence peptidase with two known substrates. One substrate is TasA (Tjalsma et al., 1998; Stover and Driks, 1999a). The other substrate is the first gene product of the operon. Up to now we have referred to that first gene product as YqxM, based on the prior unknown function designation of its gene product. Indeed, this is the protein involved in placing TasA in the matrix and mutants of which yield D-Tyr-resistant biofilms (Branda et al., 2006; Kolodkin-Gal et al., 2010). Here we show that this protein is required for the proper anchoring and polymerization of TasA at the cell surface. Furthermore, we show that much of this protein is found in discrete foci on the cell surface and that these foci disappear upon treatment with D-amino acids. In addition, we found that this protein is a minor component of the amyloid fibres and that in cells lacking it, TasA appears to be degraded. Thus, this protein has multiple functions. It serves to anchor the amyloid fibres to the cell as well as to nucleate fibre polymerization and is a component of the fibres. Therefore, we name it TapA for TasA anchoring/assembly protein and refer to it as such throughout this paper. Results TapA and TasA must be synthesized in the same cell to produce functional amyloid fibres Bacillus subtilis NCIB3610 forms floating biofilms, known as pellicles, at the liquid-air interface of standing liquid cultures (Branda et al., 2006). When fully formed, pellicles of wild-type cells are characterized by robust wrinkling (Fig. 1A). This phenotype requires the expression of two operons: epsa-o (henceforth referred to simply as eps) and tapa-sipw-tasa (formerly referred to as the yqxmsipw-tasa operon). The products of these operons form

3 B. subtilis amyloid fibre accessory protein 1157 the two major components of the biofilm matrix: the exopolysaccharide and the TasA amyloid fibres (Branda et al., 2001; 2006). The fragile pellicles of eps mutants are clearly different from the flat pellicles of tasa and tapa mutants (Fig. 1A). Previously, we showed that extracellular complementation of biofilm formation occurs when eps and tasa mutants are mixed (Branda et al., 2006). We repeated these extracellular complementation experiments and added the observation that mixing of eps and tapa mutants yielded a wild-type pellicle as well (Fig. 1B, tasa + eps and tapa + eps). However, no complementation was obtained when tapa and tasa mutants were mixed (Fig. 1B, tapa + tasa). These data suggest that the TapA and TasA proteins must be produced in the same cell in order to assemble functional fibres. An alternative possibility is that the TasA fibres from tapa mutants are not attached to cells and are therefore diluted in the pellicle medium. We therefore also assessed extracellular complementation of the tapa and tasa mutants on agar surfaces and were unable to observe complementation as determined by colony wrinkling and Congo Red binding (Fig. S1). Thus, we favour the hypothesis that the two proteins must be produced in the same cell to form functional amyloids. Punctate localization of TapA on the cell wall The finding that TapA and TasA must be produced in the same cell to yield wild-type biofilms prompted us to initiate TapA localization studies. In the study showing that different D-amino acids incorporated into the cell wall led to biofilm disassembly, we obtained mutant forms of TapA that resulted in biofilms that were resistant to D-tyrosine (Kolodkin-Gal et al., 2010). This provided initial indirect evidence that TapA was somehow associated with the cell wall. The fluorescence and electron microscopy results we present now further suggest that TapA is a cell wallassociated protein. Using anti-tapa antibodies and a FITC-conjugated secondary antibody it was possible to visualize TapA clusters in wild-type cells from pellicles after 24 h of growth, when pellicles have reached maturity. Interestingly, most of the TapA is observed as puncta located on the cell surface (Fig. 2A). Treatment of wildtype pellicles with 3 mm D-Tyr led to the disappearance of the fluorescent puncta (Fig. 2B). In a prior work, we had shown that treatment with D-Tyr does not alter transcription of the tapa operon (Kolodkin-Gal et al., 2010). Rather, D-Tyr treatment leads to the release of the fibres from the cells (Kolodkin-Gal et al., 2010). Thus it appears that loss of puncta correlates with release of fibres. Importantly, puncta remained after treatment with 3 mm D-Tyr of a strain bearing a D-Tyr-resistant mutation in tapa (Fig. 2C) and no puncta were visible in a tapa mutant (Fig. 2D). The results discussed above suggest an association of TapA with the peptidoglycan, but they are insufficient to show that the two macromolecules interact physically. However, two additional lines of evidence point to the existence of a close association between TapA and peptidoglycan. First, immunogold electron microscopic analyses revealed large concentrations of TapA at foci on the peptidoglycan (Fig. 2E). Second, cell fractionation studies localized a functional TapA YFP fusion protein to the peptidoglycan fraction and not on the protoplast fraction (Figs S2 and S3). Altogether, the results indicate that TapA is closely associated with the peptidoglycan. This led us to ask whether TapA co-purifies with peptidoglycan. TapA is associated with purified sacculi and retains functionality To obtain purified peptidoglycan sacculi, wild-type, tasa and tapa mutants were grown for 24 h in pellicle biofilms and then subjected to several washes with a solution of SDS and b-mercaptoethanol and high temperatures to dissolve the cellular content (Fig. 3). To determine whether the purified sacculi retained TapA, we analysed the samples by transmission electron microscopy (TEM) and immunogold labelling. Strikingly, despite the harsh treatment, TapA remained associated to the sacculi (Fig. 3). Importantly, only sacculi from wild-type and the tasa mutant were decorated with anti-tapa antibodies. No signal was associated to sacculi purified from a tapa mutant (Fig. 3A). We previously demonstrated that purified TasA fibres added to a tasa mutant became cell associated and restored the formation of biofilms (Romero et al., 2010). We therefore wondered whether TapA in purified sacculi would retain the ability to associate with TasA fibres. To test this, a solution of purified TasA fibres was mixed with the different sacculi and incubated for 10 h at room temperature. Excess TasA was removed by centrifugation and the samples were suspended in PBS prior to TEM and immunogold labelling with anti- TasA antibody (Fig. 3B). Electron microscopy showed that the added TasA fibres were tightly associated with sacculi from wild-type and tasa mutant cells but not with those of a tapa mutant (see Fig. 3B top panels with the bottom panels showing close-up views). Quantification of the gold-labelled particles showed that the tapa mutant sacculi bound about 10% of the fibres relative to the wild-type sacculi (Fig. S4A). As a complement to this experiment, we used western analysis with anti-tasa antibody to quantify the amount of TasA that remained associated with the sacculi in the above experiment. Using these analyses, we found the tapa mutant sacculi bound about 16 fewer TasA fibres than the wild-type sacculi (Fig. S4B). Thus, we conclude that TapA is present in the peptidoglycan where it functions as an anchor point for

4 1158 D. Romero, H. Vlamakis, R. Losick and R. Kolter Fig. 2. Localization of TapA is punctate. A D. Immunocytochemistry with anti-tapa antibody and FITC-conjugated secondary antibody performed on intact cells from 24 h pellicles grown in MSgg broth. (A) Wild-type cells show foci of TapA. (B) Wild-type cells grown in the presence of 3 mm D-Tyr. (C) Strain IKG44, a spontaneous mutant resistant to D-amino acids, grown in the presence of 3 mm D-Tyr. (D) tapa mutant cells that do not have signal. E and F. Electron micrographs of thin sections of resin-embedded wild-type cells after 24 h growth and immunogold labelling with anti-tapa antibody. (E) Wild-type. Arrow points to the accumulation of gold particles in the electron-dense cell wall. (F) tapa mutant does not show any signal. Scale bars, 2 mm in (A) (D) and 100 nm in (E) and (F). TasA fibres. Moreover, TapA retains functionality as an anchor even after isolation of sacculi. TapA is a minor component of TasA fibres in the extracellular matrix While the above experiments indicate that TapA is present in the peptidoglycan, they do not rule out the possibility that the protein might also be present in the extracellular matrix. To determine whether TapA is present in the matrix, we separated the biofilm into medium (Med), cell and matrix (Mat) fractions and performed immunoblot analyses with anti-tapa antibody. Because biofilm formation is dynamic, we analysed samples collected over time at a constant incubation temperature (30 C). Under these conditions pellicles reached maturity at 24 h and by 48 h began to disassemble. Results of these experiments are shown in Fig. 4. At 24 h, two TapA-related bands were detected in both cells (Cell) and matrix (Mat); no TapArelated proteins were detected in the medium (Med). The larger band corresponded to 28 kda, in close agreement with the predicted mass of secreted TapA, whose signal peptide has been cleaved off. The small band, corresponding to 24 kda, probably represents a further processing of TapA. Importantly, neither of the two bands was detected in the tapa mutant (Fig. 4). After 48 h of incubation, when disassembly was just beginning, only the smaller 24 kda band remained in the extracellular matrix (Mat) and no signal could be detected in the cell fraction. Thus, TapA localization varies during biofilm development.

5 B. subtilis amyloid fibre accessory protein 1159 Fig. 3. TapA is associated with the cell wall. Transmission electron micrographs and immunolabelling of B. subtilis sacculi. A. Anti-TapA antibody. Arrows point to gold particles present in wild-type and tasa mutant cells that are absent in the tapa mutant B. Fibres of TasA (+TasA) purified from B. subtilis were added to sacculi from the designated strains and immunogold labelling was performed with anti-tasa antibody. Bottom panel is a magnified view of the boxes marked in (B). Scale bars, 200 nm. In addition, it appears that TapA undergoes processing beyond the removal of the signal peptide and that this processed form of the protein remains matrix-associated at 48 h of biofilm development. A similar processed product of TapA was observed previously in shaken cultures, before there was knowledge of its involvement in the production of an extracellular matrix (Stover and Driks, 1999b). Finding TapA present in the matrix led us to ask whether, similar to EPS and TasA, TapA might also be a structural component of the matrix. We hypothesized that TapA might be incorporated into the TasA fibres similar to

6 1160 D. Romero, H. Vlamakis, R. Losick and R. Kolter Fig. 4. TapA localizes to the cell and the matrix. Biochemical fractionation of wild-type and tapa pellicles analysed by immunoblot with anti-tapa antibody. Pellicles of cells grown in MSgg medium after (A) 24 h and (B) 48 h were mildly sonicated and separated into three fractions: medium (Med), cell and matrix (Mat). Each fraction was concentrated prior to analysis by SDS-PAGE. CsgB in Curli fibres or other minor pilin proteins in pili of Gram-positive bacteria (Bian and Normark, 1997; Kline et al., 2010). To test this hypothesis, we carried out immunogold electron microscopy using gold particles of different diameters to simultaneously detect TapA and TasA in biofilm samples. Initial microscopy studies were done with strain DR4, which harbours a non-polar mutation in tapa and a functional TapA YFP translational fusion (fusion functionality is demonstrated in Fig. S2). Biofilms of this strain were analysed after 24 h of growth by TEM and co-immunogold labelling with anti-tasa and anti-yfp antibodies (Fig. 5). For these localization experiments, gold particles of 10 nm were used to detect TasA and gold particles of 15 nm were used to detect TapA YFP. Indeed, fibres were decorated with both antibodies, indicating that TapA is incorporated into the TasA fibres (Fig. 5). Negative controls were performed with the anti-yfp antibody on wild-type samples that did not harbour any YFP fusion and little to no signal was detected, indicating that the YFP antibody does not react non-specifically with the cells (data not shown). The finding of TapA in the amyloid fibres was surprising because prior analyses had suggested that the fibres were composed largely of TasA. When fibre preparations were subjected to SDS-PAGE there was only one band visible TasA after Coomassie blue staining [see Fig. S2 in (Romero et al., 2010)]. We repeated those immunoblot experiments but now with both anti-tasa and anti-tapa antibodies. TapA was only detected in Western blot analysis of purified TasA fibres after treatment with formic acid and a 200-fold concentration of the sample (see Fig. S5A). Interestingly, the size of the reacting band coincided with the molecular weight of TapA present in the matrix in fractionation experiments (see Fig. 4). We quantified the ratio of TasA to TapA in fibres purified from B. subtilis by performing quantitative immunoblot experiments using purified recombinant His 6-TasA and His 6-TapA as standards (Fig. S5B). We calculated that the ratio of TasA to TapA monomers was approximately 100:1. A tapa mutant produces fewer and altered fibres Fig. 5. TapA and TasA localize to fibres. Electron micrograph of negatively stained, immunogold-labelled samples from pellicles of strain DR4 after 24 h of incubation in MSgg medium at 30 C. Two sizes of gold particles were used to detect TasA (10 nm) or YFP (15 nm). Arrows point to different sized gold particles. Scale bar, 100 nm. The findings presented thus far show that (i) TapA and TasA must be made in the same cell to be functional, (ii) TapA accumulates as a focus in the cell wall and (iii) TapA is a minor component of the amyloid fibres. All of these data are consistent with the hypothesis that TapA might be a nucleator for TasA polymerization. To begin to explore this possibility, we examined the fate of TasA in a strain harbouring a non-polar tapa mutation. We used TEM and immunogold labelling with anti-tasa antibodies to analyse wild-type and tapa mutant biofilms after 24 h of growth. In wild-type biofilms we observed TasA forming large fibres anchored to cells (Fig. 6A). In contrast, in the tapa mutant, most cells showed anti-tasa signal near the surface in the cytosol (Fig. 6B) and only rarely did we observe a few disorganized fibrils decorated with anti- TasA antibody and these were invariably dissociated from cells (Fig. 6C). In addition, we examined the amyloid properties of the residual TasA fibrils found in the tapa mutant. When grown on solid medium containing the amyloid-

7 B. subtilis amyloid fibre accessory protein 1161 Fig. 6. TapA is required for TasA fibre formation. Electron micrographs of negatively stained, anti-tasa immunogold-labelled samples from pellicles grown for 24 h in MSgg medium. A. Wild-type cells. B and C. tapa mutant cells did not harbour TasA-labelled fibres, but instead had cell-associated TasA signal (B). In rare occasions small and disorganized fibres of TasA were observed detached from cells (C). The bottom panel represents a magnified view of areas marked in (A) (C). Scale bars, 200 nm. binding dye Congo Red, the wild-type colonies stained red whereas the mutant colonies were brown, suggesting a lack of amyloids (Fig. 7A). Next, we subjected the tapa mutant to the same 1 M NaCl extraction procedure that we routinely use to prepare TasA fibres from wild-type cells (Romero et al., 2010). We examined the extract for its ability to bind the amyloid-binding dye Congo Red and measured the absorption at four wavelengths from 400 nm to 650 nm (Fig. 7B). The extract from wild-type cells shows a characteristic peak at 540 nm. This peak was much reduced in the tapa mutant, resembling more the sample that contained Congo Red alone. Together these data suggest that while a few TasA fibres polymerize in the absence of TapA, these are thin and short, and do not appear to retain the amyloid properties characteristic of the fibres formed by wild-type cells. Furthermore, as seen in the flat pellicle in Fig. 1, the fibres produced in the tapa mutant were incapable of forming wrinkled biofilms. The amount of TasA is dramatically reduced in a tapa mutant Mutants lacking tapa produce few fibres and these do not display obvious amyloid properties (see Figs 6 and 7). It is possible that the decreased ability to bind Congo Red is simply due to a much lower total protein concentration. To determine whether the decrease in amyloid properties was due to an overall decrease in TasA protein, similar quantities of wild-type and tapa mutant pellicles were subjected to SDS-PAGE and the gels were analysed by immunoblot with anti-tasa antibody (Fig. 8A). We observed much less TasA protein in the tapa mutant compared with wild-type. As expected, the tasa mutant cell extracts did not react with the antibody. To make sure that the decrease in TasA labelling was not due to secretion of TasA in the absence of TapA, we assayed the medium, as well as the matrix and cell fractions from a pellicle assay. We only observed TasA labelling in the cell fraction suggesting that there is not excess TasA secreted from that strain (Fig. S6). A reasonable explanation for the decreased amount of TasA in a tapa mutant could be reduced transcription from the promoter of the tapa-sipwtasa operon. To test this hypothesis, we analysed wildtype and tapa mutant cells harbouring a P tapa-yfp reporter by flow cytometry (Fig. 8B). Both strains showed similar fluorescence profiles of expression at both 24 h and 48 h indicating that there is not decreased transcription of tasa in the tapa mutant. In addition, we constructed a strain in which the expression of tasa was uncoupled from the P tapa

8 1162 D. Romero, H. Vlamakis, R. Losick and R. Kolter we were unable to observe complementation when TasA fibres were added to the tapa mutant (Fig. 9B). Next, the entire complemented pellicles shown in Fig. 9A and B were harvested and cells were pelleted to remove excess fibres prior to TEM immunogold analysis with anti-tasa antibodies. We observed TasA-labelled fibres emanating from cells of the tasa mutant (Fig. 9C). However, in the tapa mutant, the purified TasA appeared as small and disorganized fibrils (Fig. 9D). Thus, the tapa mutant cells cannot bind pre-formed amyloid filaments. This observation is in agreement with the results shown in Fig. 3 in which sacculi from tapa mutant cells could not bind purified amyloid fibres. Discussion Fig. 7. tapa mutant cells and extracts do not bind Congo Red. A. Colonies grown for 72 h on Congo Red indicator plates. B. Absorbance of a 10 mm solution of Congo Red incubated alone or with extracts from wild-type or tapa mutant cells. promoter and instead was placed under the control of the IPTG-inducible P hyperspank promoter. Induction of tasa transcription by the addition of 0.2 mm IPTG was enough to complement a tasa mutant biofilm morphology; however, this was not sufficient to complement a tapa mutant (Fig. S7). These findings taken together suggest that in the absence of TapA, the TasA protein does not form amyloid fibres and is thus less stable and may be a substrate for proteolytic degradation (Tjalsma et al., 2004). Amyloid fibres are produced by bacteria to serve as a major structural component of the extracellular matrix that holds biofilm-associated cells together (Collinson et al., 1993; Chapman et al., 2002; Claessen et al., 2003; Dueholm et al., 2010; Romero et al., 2010). In B. subtilis biofilms the amyloid fibres are primarily composed the TasA protein, which is encoded in the tapa-sipw-tasa operon. While we had studied TasA in some detail, the precise function of TapA was not clear. We had previously obtained evidence suggesting that TapA was involved in getting TasA to the matrix (Branda et al., 2006). Recently, we found that at late stages in the life cycle of a biofilm, specific D-amino acids, including D-tyrosine, are produced and incorporated in the cell wall (Kolodkin-Gal et al., Purified amyloid fibres cannot complement a tapa mutant extracellularly Given that the tapa mutant is defective in TasA fibre production, we needed to re-think the experiment shown in Fig. 1 where the tapa and tasa mutants were unable to complement each other extracellularly. This inability could simply be due to the fact that both mutants are unable to produce functional TasA amyloids. Thus, we asked whether purified TasA amyloid fibres could restore biofilm formation to a tapa mutant. Using the same experimental approach we reported previously, we reproduced the ability of purified TasA fibres to restore pellicle formation to a tasa mutant (Romero et al., 2010) (Fig. 9A). However, Fig. 8. TasA protein levels are low in a tapa mutant. A. Immunoblot with anti-tasa antibodies of whole cells obtained from 24 h or 48 h MSgg pellicles from wild-type, tapa and tasa mutant strains. B. Flow cytometry analysis showed no variations in the dynamics of P tapa-yfp expression in wild-type (black line) and tapa mutant cells (red line). Grey peak corresponds to background of cells with no fluorescent reporter. Y-axis is number of cells and x-axis is fluorescence intensity in arbitrary units.

9 B. subtilis amyloid fibre accessory protein 1163 Fig. 9. Purified TasA fibres cannot restore pellicle formation to tapa mutant cells. A and B. Pellicles grown in MOLP broth for 48 h. Purified TasA fibres were included in the +TasA panels at the beginning of the experiment. (A) tasa mutant. (B) tapa mutant. C and D. Anti-TasA immunogold-labelled electron micrographs of cells from the +TasA panels. (C) TasA fibres are present in the tasa mutant + TasA pellicle and (D) absence of signal in the tapa mutant after addition of purified TasA. Scale bars 500 nm. 2010). This results in biofilm disassembly due to a release of the cells from the TasA fibres. Interestingly, mutants in the C-terminal domain of TapA confer resistance to the biofilm-dissociating activity of D-tyrosine. This gave rise to the hypothesis that TapA might function to anchor TasA fibres to the cell wall. In this paper we tested this hypothesis and showed that, indeed, TapA serves as an anchor and assembly factor for TasA fibres. We found that TapA localizes to the cell wall where it is necessary to support TasA fibre formation and attachment. Furthermore, in the absence of TapA, TasA appears to be degraded. Finally, in addition to its cell wall localization, TapA is also present as a minor component of the TasA fibres. Genetic, fractionation and microscopy experiments all indicate that TapA is associated with the cell wall. Interestingly, TapA remained associated with the peptidoglycan in sacculi preparations. This procedure involves numerous washes in 1% SDS, 0.5% beta mercaptoethanol, 2 M NaCl and treatment at 80 C. Because of its ability to stay associated with peptidoglycan through this harsh treatment, we suspect TapA is covalently linked to the cell wall. One way that proteins become covalently linked to the cell wall in Gram-positive bacteria is via sortase activity. Sortases are enzymes that recognize C-terminal sorting sequences, frequently an LPXTG motif, and catalyse attachment of these secreted proteins to the peptidoglycan (Tjalsma et al., 2004; Marraffini et al., 2006). However, TapA does not have a sequence that is similar to a known sortase domain. Nonetheless, we tested whether TapA functionality depended on the two predicted sortases of B. subtilis, the products of the yhcs and ywpe genes (Tjalsma et al., 2000a), by mutating both of these genes. The double mutant was not defective in biofilm formation, suggesting they are not involved in cell wall anchoring of TapA (data not shown). How TapA is linked to the cell wall, and whether it is through a covalent bond are questions that are currently under investigation. In addition to their role in linking proteins to the cell wall, some sortases have also been shown to function in assembling of pili in Gram-positive bacteria (Marraffini

10 1164 D. Romero, H. Vlamakis, R. Losick and R. Kolter et al., 2006). In B. cereus, pili are composed of a major subunit, BcpA, and a minor subunit BcpB. BcpA harbours an LPXTG motif that is recognized by Sortase A and allows covalent linkage to the cell wall (Budzik et al., 2008). In addition, BcpA has a YPKN motif that is recognized by a different sortase, Sortase D, which allows for BcpA to be linked to BcpB (Budzik et al., 2009). Of note, TasA harbours a YPKN motif in its sequence that tempted us to speculate a similar mechanism might be occurring between TasA and TapA during fibre assembly in B. subtilis. However, as stated above, even a double mutant in the two predicted sortase genes of B. subtilis did not alter biofilm morphology. Once again, we are left with an elusive mechanism of assembly. TasA is an example of a growing number of bacterial proteins that form surface structures with amyloid properties. These amyloid-like fibres have now been described in diverse species such as E. coli, Salmonella enterica, S. coelicolor, Pseudomonas fluorescens and B. subtilis (Collinson et al., 1993; Chapman et al., 2002; Claessen et al., 2003; Dueholm et al., 2010; Romero et al., 2010). There is vast diversity in primary sequence of these amyloidogenic proteins and they are found in Gram-positive and Gram-negative organisms, which have dramatic differences in their cell wall structures. Thus, it is likely that there are also major differences in the mechanisms of fibre assembly and attachment to cells in the different organisms. Fibres of TasA appear to polymerize in vitro following a kinetics that matches a nucleation-polymerization model, described previously for curli in E. coli (Naiki and Gejyo, 1999; Romero et al., 2010). Similar to the protein subunits of curli, CsgA and CsgB, we hypothesized that polymerization of TasA on cell surfaces could be promoted by TapA. In the absence of TapA, we observed some auto-assembly of TasA, but only in small fibres that were detached from cell. This is different from curli assembly in E. coli, where in the absence of CsgB there is no fibre formation (Barnhart and Chapman, 2006). Our data also indicate that TasA is less stable in the absence of TapA, suggesting that TapA might play a role in stabilizing TasA. This is similar to what is observed in E. coli curli, where an accessory factor, CsgG, has been shown to stabilize CsgA and CsgB, the subunits composing the amyloid fibres (Hammar et al., 1996). What is the biological function of TapA? TapA accumulates in the cell wall in young biofilms, when active polymerization of TasA fibres is needed. At this point, TapA appears to be involved in the formation and anchoring of TasA fibres. Perhaps TasA does not polymerize well in the absence of TapA and is thus rendered unstable. As the biofilms age it may become advantageous for cells to dissociate from the network of amyloid fibres. Because of the intrinsic stability of amyloid fibres, it is possible that depolymerizing the fibres is an inefficient process at best. Therefore, the Achilles Heel of the matrix may be the point where the fibres are anchored to the cell envelope. Maintaining an amyloid anchoring protein such as TapA provides an ideal mechanism for eventual disassembly of the biofilm. There is no need to break down the amyloid fibres. Instead, they are simply released when they are not required anymore. To do this, D-amino acids are produced and incorporated into the cell wall. This disrupts the anchoring of TapA, and cells are freed from the network of amyloid fibres. We propose that at the time of biofilm disassembly, the role of TapA is to provide a weak link that makes disassembly far simpler than destruction of the amyloid fibres. Thus, TapA is a bifunctional protein that allows for both assembly and detachment of the TasA amyloid fibres during biofilm formation. It will be interesting if other amyloid fibre anchoring proteins have properties similar to TapA. Experimental procedures Growth media and culture conditions Luria Bertani (LB) broth: 1% tryptone (Difco), 0.5% yeast extract (Difco), 0.5% NaCl. MSgg broth: 100 mm morpholinepropane sulphonic acid (MOPS) (ph 7), 0.5% glycerol, 0.5% glutamate, 5 mm potassium phosphate (ph 7), 50 mg ml -1 tryptophan, 50 mg ml -1 phenylalanine, 2 mm MgCl 2, 700 mm CaCl 2,50mM FeCl 3,50mM MnCl 2,2mM thiamine, 1 mm ZnCl 2 (Branda et al., 2001). MOLP broth as described previously (Romero et al., 2010). Media were solidified with the addition of 1.5% agar. For colony architecture, 3 ml of starting culture was spotted onto MSgg agar plates and incubated at 30 C for the indicated times. For pellicle formation, 12 ml of starting culture was added to 4 ml or 1 ml of MSgg or MOLP broth in a 6- or 24-well microtitre dish, respectively, and incubated without agitation at 30 C for the indicated time. For Congo Red binding assay, MSgg plates containing 20 mg ml -1 Congo Red and 10 mg ml -1 Coomassie brilliant blue G were spotted with cells as described above and assayed after 72 h at 30 C (Romero et al., 2010). For extracellular complementation, starting cultures were mixed with 40 mg of purified protein and the mixtures were used to form pellicles, as described above. Antibiotic concentrations (final): MLS (1 mg ml -1 erythromycin, 25 mg ml -1 lincomycin); spectinomycin (100 mg ml -1 ); tetracycline (10 mg ml -1 ); chloramphenicol (5 mg ml -1 ); kanamycin (10 mg ml -1 ). Strain construction Strains used and generated in this study are listed in Table 1. Plasmids were constructed and amplified in E. coli XL-1 Blue (Strategene) following manufacturer protocols. To construct the plasmid pdfr2 (laca::p tapa-tapa-yfp, spc) a 1250 base pair DNA fragment containing the promoter sequence of tapa and the open reading frame of the tapa gene, without stop codon, was amplified using the primers P tapa-f (5 - tggcgaattctcagagttaaatggtattgcttcact-3 ) and tapa-r (5 -

11 B. subtilis amyloid fibre accessory protein 1165 Table 1. Strains used in this study. Strain Genotype Reference B. subtilis 168 Prototroph Branda et al. (2001) SSB , (tapa-sipw-tasa)::spc Branda et al. (2004) NCIB 3610 Wild-type. Undomesticated strain Branda et al. (2001) SSB488 eps::tet Branda et al. (2006) CA017 tasa::km Vlamakis et al. (2008) FC268 (tapa-sipw-tasa)::spc Chu et al. (2006) amye::(tapa(13 234)-sipW-tasA) (cm) ZK3755 laca::p tapa-yfp Lopez et al. (2009) IKG44 NCIB 3610 TapA6 Kolodkin-Gal et al. (2010) DR4 (tapa-sipw-tasa)::spc This study amye::(tapa(13 234)-sipW-tasA) (cm), laca::p tapa-tapa-yfp DR5 (tapa-sipw-tasa)::spc This study amye::(tapa(13 234)-sipW-tasA) (cm), laca::p hyperspank-tasa DR6 tasa::km, laca::p hyperspank-tasa This study DR7 (tapa-sipw-tasa)::spc This study amye::(tapa(13 234)-sipW-tasA) (cm), laca::p tapa-yfp ctgatcagcttcattgcttttttcatc-3 ). A fragment of 600 base pair containing the whole sequence of yfp was amplified from plasmid pkm003 using the primers yfp-f (5 -gatgaaaaa agcaatgaagctgatcagatgagtaaaggagaagaactt-3 ) and yfp-r (5 -gcgctcaggatccttatttgta-3 ). Long flanking homology PCR was used to join the PCR fragments. The resulting PCR product was digested with EcoRI and BamHI enzymes and cloned into the plasmid pdr183 (Doan et al., 2005), cut with the same enzymes. The plasmid pdfr4 (laca::p hyperspank-tasa) was constructed in two steps. First, the open reading frame of tasa was amplified using the primers TasA-F (5 -aataaaagtcgacat aaaaggggagcttaccatgggtatgaa-3 ) and TasA-R (5 -tttgcat GCttattaatttttatcctcgctatgcgc-3 ). The PCR product was digested with SalI and SphI enzymes and cloned into the plasmid pdr111 cut with the same enzymes to generate the plasmid pdfr3 (amye::p hyperspank-tasa). Then pdfr3 was digested with XhoI and BamHI and the fragment containing P hyperspank-tasa was subcloned into the plasmid pdr183 digested with the same enzymes, resulting in pdfr4 (laca::p hyperspank-tasa). To construct the plasmids pdfr5 (pet22b-tapa) and pdfr6 (pet22b-tasa), a fragment of DNA containing the open reading frame without signal peptide or stop codon of the tapa or tasa genes was amplified by using the primers TapAH-F (5 -aaaaaaaaa-catatgatatgcttacaatttttc- 3 ) and TapAH-R (5 -aaaaaaaaa-ctcgagctgatcagcttcatt gct-3 ) or TasAH-F (5 -aaaaaaaaa-catatggcatttaacgacatta aa-3 ) and TasAH-R (5 -aaaaaaa-ctcgagatttttatcctcgctat gcgc-3 ) respectively. Both fragments were digested with NdeI and XhoI enzymes and cloned into the plasmid pet22-b, cut with the same enzymes. To generate the strains DR4 ((tapa-sipw-tasa)::spc, amye::(tapa(13 234)-sipW-tasA) (cm), laca::p tapa-tapa-yfp, mls) and DR5 ((tapa-sipw-tasa)::spc, amye::(tapa(13 234)- sipw-tasa) (cm), laca::p hyperspank-tasa, mls), the mutant strain lacking the tapa-sipw-tasa operon (SSB149) was transformed by natural competence with pdfr2 and pdfr4, respectively, and positive clones were used as donor strains for transferring the constructs into B. subtilis strain mutant (FC268) by means of SPP1-mediated generalized transduction (Yasbin and Young, 1974). To generate DR6 (tasa::km, laca:: P hyperspank-tasa, mls), B. subtilis strain 168 was transformed with plasmid pdfr4 and subsequently transduced into CA017. For construction of the strain DR7, the strain ZK3755 [B. subtilis 3610 containing the translational fusion laca::p tapa-yfp (Lopez et al., 2009)] was used as donor and transduced into strain FC268 (Chu et al., 2006). Protein purification TasA produced by B. subtilis was purified as previously described (Romero et al., 2010). Briefly, cells were grown in MSgg broth at 30 C for 20 h. After centrifugation, the pellet containing cells was extracted twice with saline extraction buffer supplemented with a protease inhibitor mixture (Complete; Roche). Supernatant containing the protein was separated from cells by centrifugation and filtered through a 0.4 mm polyethersulfone bottle-top filter. This procedure purified fibres containing TasA and the product was stored at -20 C prior to use. The plasmids pdfr5 and pdfr6 were used for the production of His6-TapA and His6-TasA fusion proteins respectively. These plasmids were transformed in BL21 competent E. coli cells. Cultures in 100 ml of LB supplemented with Amp were grown shaking at 37 C to an OD 600 of 0.5. IPTG was added to a final concentration of 1 mm and the cultures were incubated for additional 3 h. Cells were harvested by centrifugation, resuspended in 15 ml of lysis buffer, 1 CelLytic B cell lysis reagent (Sigma) diluted in 20 mm Tris, 500 mm NaCl, 20 mm Imidazol and 1 mm PMSF, and supplemented with 100 mg ml -1 of freshly prepared lysozyme solution, and incubated for 15 min. Further disruption and reduction of viscosity was done by sonication. Approximately 1.5 ml of nickel chelating resin (G- Bioscience) was washed and conditioned as indicated by the manufacturer prior to adding to the samples and incubating with gentle agitation for 1 h at room temperature. The mixture was centrifuged and after decanting the supernatant, the lysate/resin mixture was washed with 5 volumes of binding buffer (20 mm Tris, 500 mm NaCl, 20 mm Imidazol, 1 mm

12 1166 D. Romero, H. Vlamakis, R. Losick and R. Kolter PMSF), 3 volumes of washing buffer-i (20 mm Tris, 500 mm NaCl, 40 mm Imidazol, 1 mm PMSF) and 3 volumes of washing buffer-ii (20 mm Tris, 500 mm NaCl, 100 mm Imidazol, 1 mm PMSF). The proteins were eluted with elution buffer (20 mm Tris, 500 mm NaCl, 500 mm Imidazol, 1 mm PMSF). Purified proteins were stored at -20 until used. Cell fractionation assay Cell fractionation assays were performed as described previously (Tjalsma et al., 2000b). Cells grown in biofilm conditions were separated from the extracellular matrix by sonication as described previously (Branda et al., 2006). Protoplasts were obtained from by treatment of cells with 0.5 mg ml -1 lysozyme in protoplast buffer (Tjalsma et al., 2000b) for 30 min. Protoplast supernatant containing cell wall-associated proteins was separated from protoplasts by mild centrifugation, and the protoplasts were resuspended in fresh protoplast buffer. A fraction of the protoplast preparation was additionally treated with 1% Triton X-100 to release cytoplasmic proteins and spores. Immunoblot and quantitative immunoblot Samples were analysed by SDS-PAGE with either 12% or 10% polyacrylamide and blotted onto PVDF membrane using standard protocols. For detection of TasA, blots were probed with anti-tasa antibodies raised in rabbits used at dilution of 1: In the fractionation assays, blots were probed with anti-yfp polyclonal antibody (Clontech) at dilution 1:5000 or anti-sigmaa antibody (a gift from D. Rudner, Harvard Medical School, Boston, MA) as a control for detecting cytoplasmic content. A secondary anti-rabbit IgG antibody conjugated to horseradish peroxidase (Bio-Rad) was used at a dilution of 1: Blots were developed using the Pierce super signal detection system (Pierce, Thermo Scientific). TapA antibody was kindly provided by A. Driks, Loyola University Medical Center, Maywood, IL. Quantification of TasA and TapA proteins in purified fibres was done by quantitative immunoblot as previously described (Chai et al., 2008). The ratio TasA : TapA was estimated to be 100:1 by comparing the pixel density of Fig. S3B fibres from B. subtilis to known amounts of purified proteins from E. coli. In order to visualize the TapA, fibres were concentrated 20-fold for the anti-tapa western relative to the anti-tasa blot. room temperature. Then the sacculi were recovered by soft centrifugation, resuspended in a minimal volume of PBS and analysed by TEM and immunogold labelling with anti-tasa antibody. TEM and immunolabelling Observation of intact cells was done from 24 h pellicles. Samples were adsorbed onto carbon or formvar/carbon coated grids, which were previously treated to make the grids hydrophilic. The excess sample was blotted off using filter paper and the grids were floated on 5 ml of 1 2% uranyl acetate for a few minutes. The samples were dried prior to examination. For immunolocalization studies, samples were floated on blocking buffer (1% non-fat dry milk in PBS with 0.1% Tween 20) for 30 min and on anti-tasa or anti-yfp primary antibodies diluted 1:150 in blocking buffer for 2 h, rinsed in PBST and exposed to goat-anti-rabbit 20 nm gold secondary antibody diluted 1:50 (TedPella) for 1 h, and rinsed. For double labelling, samples were fixed in 1% glutaraldehyde for followed by quenching in 4 drops of 0.15 M glycine/pbs prior to proceeding with primary and secondary antibodies conjugated to different sized gold particles as indicated in the text. All grids were stained with uranyl acetate and lead citrate for visualization. For subcellular localization studies of TapA, cells grown for 24 h in pellicle assays were separated by mild sonication as previously described. Cell suspensions were fixed with an equal volume of 4% paraformaldehyde in 0.1 M sodium phosphate buffer, ph 7.4. Small pieces of cell pellet were infiltrated in 2.3 M sucrose in PBS containing 0.15 M glycine. The samples were high pressure frozen in a Leica EM Pact2 high-pressure freezer and freeze-substituted in 0.2% glutaraldehyde + 0.1% uranyl acetate in Acetone at -90 C for 72 h. Samples were slowly warmed up to room temperature and embedded in LR White resin. Ultrathin sections were cut at -120 C with a cryo-diamond knife. Sections were transferred to a formvar/carbon coated copper grid and stored at 4 C until immunogold-labelled. For contrasting, grids were floated in mixture of 2% methyl cellulose with 3% uranyl acetate for 10 min. Excess liquid was removed and grids were allowed to dry before examining in a Tecnai G2 Spirit BioTWIN microscope at an accelerating voltage of 80 KV. Images were taken with an AMT 2k CCD camera. Preparation of sacculi Sacculi from the B. subtilis strains NCIB 3610 (wild-type), tasa and tapa were obtained as previously described (Reusch, 1982). Briefly, cells were grown in MSgg broth standing liquid cultures for 24 h. Bacteria where harvested by centrifugation and the pellets were treated five times for 15 min per treatment with extraction buffer (1% SDS and 0.5% beta mercaptoethanol) at 80 C. The remaining pellet was washed in distilled water and 2 M NaCl, for a total of 12 rinses. The obtained sacculi were stored at -20 C prior to use. For mixing experiments, a solution of TasA fibres (in buffer Tris 20 mm NaCl 50 mm, ph 7) purified from B. subtilis were mixed with different sacculi and incubated for 10 h at Immunocytochemistry, image capture and analysis Biofilms were harvested at 24 or 48 h, resuspended in 1 ml PBS buffer and dispersed with three pulses of mild sonication. Cells were separated by centrifugation and fixed with a solution of 4% paraformaldehyde for 7 min, washed in PBS and resuspended in GTE buffer (50 mm Glucose, 10 mm EDTA ph 8, 20 mm Tris-HCl ph 8) (Vlamakis et al., 2008). The fixed cells were blocked with 2% BSA/PBS for 30 min at room temperature. Primary antibody, anti-tapa or anti-yfp was used at a dilution of 1:50 in 2% BSA/PBS for 1 h at room temperature. Samples were washed 10 with PBS and exposed to the secondary goat anti-rabbit conjugated to FITC antibody (Invitrogen, Molecular Probes) at a

13 B. subtilis amyloid fibre accessory protein 1167 dilution of 1:100 in 2% BSA/PBS for 1 h at room temperature in the dark. Finally samples were washed 8 with PBS and resuspended in fresh GTE buffer before visualization. Fluorescence microscopy images were taken on a Nikon Eclipse TE2000-U microscope equipped with an X-cite 120 illumination system, using a Hamamatsu digital camera model ORCA-ER. Fluorescence signal was detected with a Ex436/500 filter. Image processing was performed using MetaMorph Software and Photoshop. Flow cytometry Biofilms of cells harbouring P tapa-yfp fusions (ZK3755 and DR7) were harvested at 24 or 48 h and dispersed as previously described. Cells were fixed in 4% paraformaldehyde solution for 7 min, washed with PBS and resuspended in GTE buffer. After fixation, single cells were obtained by mild sonication of the samples (Branda et al., 2006). For flow cytometric analysis, dilution of 1:100 of cell suspensions were measured on a BD LSR II flow cytometer (BD Biosciences) using a solid state laser. For YFP fluorescence, a laser excitation of 488 nm coupled with 530/30 and 505LP sequential filters were used. The photomultiplier voltage was set at V. Every sample was analysed by counting events using FACS Diva (BD Biosciences) software to capture the data (Lopez et al., 2009). Further analysis was performed using FlowJo software ( Acknowledgements We would like to thank members of the Kolter and Losick laboratories for helpful discussions. We thank Adam Driks (Loyola University Medical Center, Maywood, IL) for kindly providing antibodies against TasA and TapA (YqxM). We thank M. Ericsson, L. Trakimas and E. Benecchi for help and guidance in the electron microscope. This work was funded by National Institutes of Health grants to R.K. (GM58213) and R.L. 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