Bifunctionality of a biofilm matrix protein controlled by redox state

Significance

The biofilm matrix is a critical target in the hunt for novel strategies to destabilize or stabilize biofilms. Knowledge of the processes controlling matrix assembly is therefore an essential prerequisite to exploitation. Here, we highlight that the complexity of the biofilm matrix is even higher than anticipated, with one matrix component making two independent functional contributions to the community. The influence the protein exerts is dependent on the local environmental properties, providing another dimension to consider during analysis. These findings add to the evidence that bacteria can evolve multifunctional uses for the extracellular matrix components.

Abstract

Biofilms are communities of microbial cells that are encapsulated within a self-produced polymeric matrix. The matrix is critical to the success of biofilms in diverse habitats; however, many details of the composition, structure, and function remain enigmatic. Biofilms formed by the Gram-positive bacterium Bacillus subtilis depend on the production of the secreted film-forming protein BslA. Here, we show that a gradient of electron acceptor availability through the depth of the biofilm gives rise to two distinct functional roles for BslA and that these roles can be genetically separated through targeted amino acid substitutions. We establish that monomeric BslA is necessary and sufficient to give rise to complex biofilm architecture, whereas dimerization of BslA is required to render the community hydrophobic. Dimerization of BslA, mediated by disulfide bond formation, depends on two conserved cysteine residues located in the C-terminal region. Our findings demonstrate that bacteria have evolved multiple uses for limited elements in the matrix, allowing for alternative responses in a complex, changing environment.

Biofilms are assemblies of microbial cells that are attached to a surface or each other (1). Assembly is facilitated by manufacture of an extracellular matrix, which provides both structural and biochemical support to the biofilm community (2). After analysis of many diverse species, the biofilm matrix has been found to mainly comprise exopolysaccharides, extracellular DNA, and proteins, many of which form higher-order structures, such as fibers or filaments (3). Currently, there is increasing knowledge of the nature and form of the individual components in the matrix (3), but how the matrix components are deployed in the biofilm, how they interact with other elements in the matrix, and how the local physicochemical environment impacts the properties of the materials used in the biofilm are underexplored.

Bacillus subtilis is a Gram-positive, spore-forming bacterium, which is found in great abundance in the soil (4) and forms biofilms with a distinctive complex architecture (5). The biofilms display an unusual property: They are highly hydrophobic and remain as such after contact with water, inorganic or organic solvents, and commercial biocides (6). The hydrophobicity of the biofilm has been attributed to a small secreted protein named BslA (7, 8), which works alongside the fibrous protein TasA (9, 10) and the extracellular polysaccharide produced by the products of the epsA-O operon (5) to allow morphogenesis of the biofilm. Partial evidence of the mechanism used by BslA to confer hydrophobicity to the B. subtilis biofilm has been revealed (8, 11–13). Although transcription of bslA occurs uniformly within the population, secreted BslA is localized to the periphery of the biofilm, where it forms a hydrophobic layer at the air–biofilm surface (8, 12). Consistent with this finding, biophysical analysis of recombinant BslA in vitro demonstrated spontaneous formation of a stable elastic protein film at an air–water interface (12). Structural analysis revealed that BslA is an amphipathic protein consisting of an immunoglobin G-like scaffold appended with a hydrophobic “cap” (12) that can present in two forms: “cap in” and “cap out.” This property either shields (cap in) or reveals (cap out) the hydrophobic domain in response to the local environment (11). Thus, when BslA is in the aqueous environment of the matrix, it is likely that the hydrophobic cap is “hidden” toward the interior of the protein. In contrast, when BslA reaches a hydrophobic interface (e.g., air at the surface of the biofilm), it undergoes a limited conformational change to expose the hydrophobic cap. Nonetheless, the mechanism by which the BslA biofilm surface layer assembles in vivo is undefined.

B. subtilis encodes a monomeric BslA paralogue called YweA (14). Deletion of yweA does not impact the overall morphology or hydrophobicity of the biofilm (8); however, deletion in combination with removal of bslA exacerbates the biofilm defect of the single bslA deletion (8, 14). Contrary to the marginal contribution of YweA to biofilm formation, but consistent with the high level of amino acid sequence similarity, in vitro recombinant YweA undergoes the partial structural rearrangement at an interface to reveal the hydrophobic cap and forms an elastic protein film, albeit with limited stability (14). One notable difference between the primary amino acid sequences of BslA and YweA is that the BslA-like variants possess a short C-terminal extension that contains two conserved cysteine residues in a “CxC” configuration. Cysteine residues play an important role in the function of diverse proteins in a wide range of cellular processes (15), and therefore we evaluated whether the BslA CxC motif played a significant biological role or was functionally redundant. Our analysis indicates that in addition to BslA having two structural forms (cap in/cap out) (11), it also has two functional forms mediated not by the hydrophobic cap region, but by the cysteines at the C terminus: Monomeric BslA dictates biofilm structure, whereas surface hydrophobicity requires at least dimeric protein. Thus, we show that the cysteine residues are crucial for full BslA function and surmise that, in the native biofilm, electron acceptor availability influences BslA oligomerization: In the oxygen-rich, surface-exposed region, the predominant form is a dimer, whereas the anoxic environment in the depths of the biofilm prevents BslA dimerization and allows for nutrient uptake. This work gives an example of a biofilm matrix protein with two functions that can be genetically separated and where the activity is controlled by redox state.

Results

BslA Disulfide Bond Formation.

Recombinant BslA_42–181 forms a mixture of predominantly monomeric and dimeric protein in vitro, with a small amount of tetramer observed (12) (Fig. S1). The requirement of the cysteine residues (C178 and C180) for oligomerization of BslA was assessed in vitro by using purified recombinant protein where the cysteine residue(s) were either replaced with alanine or where the C-terminal 10 amino acids were removed (Fig. S1). Replacement of either C178 or C180 with alanine abolished tetrameric BslA (Fig. S1 A and C), but dimers, which could be disrupted by the addition of β-mercaptoethanol, still formed (Fig. S1). In contrast, when both cysteines were replaced with alanine, or the C-terminal 10 amino acids were deleted, the protein was restricted to a monomeric form (Fig. S1). Secondary structure analysis of the proteins by circular dichroism (CD) spectroscopy indicated that each of the BslA variants had the same overall fold as the wild-type protein, indicating that the inability to form dimers or tetramers was not due to a disruption of the protein structure (Fig. S2A).

Fig. S1.

In vitro analysis of recombinant BslA higher-order form. (A and B) SDS/PAGE analysis of 30 μg of BslA, BslA_C178A, BslA_C180A, BslA_{C178A C180A}, and BslA_Δ171–181 recombinant protein in the absence (A) and presence (B) of reducing agent. (C) SEC analysis of recombinant protein using a Superdex 75 10/300GL column in an unreduced state.

Fig. S2.

Presence of CxC motif does not affect in vitro characteristics of BslA. (A) Solution state CD spectra of recombinant BslA (black), BslA_C178A (green), BslA_C180A (blue), BslA_C178A,C180A (red), and BslA_Δ171–181 (orange) indicate all three variants have a similar secondary structure. (B) Wrinkle relaxation of WT BslA (black), BslA_C178A,C180A (red), and BslA_Δ171–181 (blue). Wrinkles were formed in the protein film by removing fluid from the droplet, and relaxation of the wrinkles was followed over time. The error bars represent the SD in normalized greyscale over at least five independent wrinkles. (C–E) Over 10 min, wrinkles did not relax noticeably. TEM images of BslA (C), BslA_C178A,C180A (D), and BslA_Δ171–181 (E) stained with uranyl acetate indicate that all three variants are capable of forming the ordered 2D lattice observed previously for WT BslA (11, 12).

Next, we tested to see whether BslA formed dimers and/or tetramers in vivo, to explore the possibility that the conserved CxC motif at the C terminus of BslA plays an in vivo role in BslA oligomerization. Proteins were extracted from the wild-type biofilm either in the presence of Cu(II)-(o-phenanthroline)₃, which cross-links disulfide bonds (16), or 10 mM DTT, a reducing agent. Western blotting revealed immunoreactive bands consistent with monomeric (∼14 kDa), dimeric (∼28 kDa), and tetrameric (∼56 kDa) forms of BslA in the absence of reducing agent (Fig. 1A), whereas in the presence of 10 mM DTT, only monomeric BslA was detected (Fig. 1B). The bslA deletion strain (NRS2097) was used as a control for antibody specificity and did not reveal any interacting bands. We generated a series of strains to express the variant bslA coding regions in the B. subtilis bslA deletion strain (Table S1). Analysis of proteins extracted from the mature biofilms by immunoblot showed the same BslA monomer, dimer, and tetramer pattern in vivo as that observed in vitro for the recombinant proteins (compare Fig. S1 with Fig. 1A). Strains generating C178A (NRS5177) and C180A (NRS5178) variant BslA formed dimers, but not tetramers, whereas the double C178A, C180A mutant (NRS5179) and the BslA_Δ172–181 (NRS2957) truncation variants were restricted to the monomeric state (Fig. 1A). In each case, only monomeric BslA was detected by Western blot analysis when 10 mM DTT was added during protein extraction (Fig. 1B). These data demonstrate that BslA forms dimers and tetramers in vivo in a C178- and C180-dependent manner.

Fig. 1.

BslA is a bifunctional protein. (A and B) Western blot analysis of BslA in a native (A) and reduced (B) state using proteins extracted from biofilms (SI Materials and Methods). (C–L) Architecture and hydrophobicity of biofilms. (C–H) Strains used were 3610 (wild-type; NCIB3610) (C), bslA⁻ (NRS2097) (D), and the bslA mutant genetically complemented with the following variants: CxC (NRS2299) (E), AxC (NR5177) (F), CxA (NRS5178) (G), AxA (NRS5179) (H), and ΔCterm (NRS2957) (I). Biofilms depicted in J–L are the result of coculturing strains NRS2299 and NRS5179. Under each biofilm is an image of a water droplet on the upper surface of the biofilm, and the values of the contact angle are in Table S2.

Table S1.

Full list of strains used in this study

Strain	Relevant genotype/description*	Source/construction†
B. subtilis
NCIB3610	Prototroph	BGSC
168	trpC2	BGSC
NRS2097	NCIB3610 bslA::cml	Ref. 45
NRS2299	NCIB3610 bslA:: cml amyE::Phy-spank-bslA-lacI (spc)	Ref. 45
NRS5165	168 amyE::Phy-spank-bslAC178A-lacI (spc)	pNW1500 → 168
NRS5166	168 amyE::Phy-spank-bslAC180A-lacI (spc)	pNW1501 → 168
NRS5167	168 amyE::Phy-spank-bslAC178AC178A-lacI (spc)	pNW1502 → 168
NRS5177	NCIB3610 bslA:: cml amyE::Phy-spank-bslAC178A-lacI (spc)	SPP1 NRS5165 → NRS2097
NRS5178	NCIB3610 bslA:: cml amyE::Phy-spank-bslAC180A-lacI (spc)	SPP1 NRS5166 → NRS2097
NRS5179	NCIB3610 bslA:: cml amyE::Phy-spank-bslA C178AC180A-lacI (spc)	SPP1 NRS5167 → NRS2097
NRS2953	168 amyE::Phy-spank-bslA Δ172–181-lacI (spc)	pNW621 → 168
NRS2957	NCIB3610 bslA:: cml amyE::Phy-spank-bslA Δ172–181-lacI (spc)	SPP1 NRS2953 → NRS2097
NRS1471	168 sacA-Pspachy-gfp (kan)	Ref. 12
NRS1473	NCIB3610 sacA-Pspachy-gfp (kan)	Ref. 12
NRS3812	NCIB3610 bslA::cat, sacA::Pspachy-gfp (kan)	Ref. 12
NRS2404	168 yweA::Kan	Ref. 44
NRS2405	NCIB3610 yweA::Kan	Ref. 44
NRS2409	168 amyE::Phy-spank-yweA (spc)	Ref. 44
NRS2412	NCIB3610 bslA:: cml amyE::Phy-spank-yweA (spc)	Ref. 44
NRS5550	168 amyE::Phy-spank-bslAss-yweA (spc)	pNW1719 → 168
NRS5551	NCIB3610 bslA::cml amyE::Phy-spank-bslAss-yweA (spc)	SPP1 NRS5550 → NRS2097
NRS4831	168 amyE::Phy-spank-bslAss-yweA-bslA171–181 (spc)	pNW1079 → 168
NRS4834	NCIB3610 bslA::cml + amyE::Phy-spank-bslAss- yweA-bslA171–181 (spc)	SPP1 NRS4831 → NRS2097
NRS5208	168 amyE::Phy-spank-bslAss-yweA-bslA171–181 C178AC180A	pNW1507 → 168
NRS5210	NCIB3610 bslA::cml + amyE::Phy-spank-bslAss-yweA-bslA171–181 C178AC180A	SPP1 NRS5208→ NRS2097
NRS5509	168 amyE::Phy-spank-bslAss-yweA-bslA171–181 C178A	pNW1702 → 168
NRS5515	NCIB3610 bslA::cml + amyE::Phy-spank-bslAss-yweA-bslA171–181 C178A	SPP1 NRS5509 → NRS2097
NRS5510	168 amyE::Phy-spank-bslAss-yweA-bslA171–181 C180A	pNW1703→ 168
NRS5516	NCIB3610 bslA::cml + amyE::Phy-spank-bslAss-yweA-bslA171–181 C180A	SPP1 NRS5510→ NRS2097
BKEE21460	168 bdbA::erm	BGSC
NRS5552	NCIB3610 bdbA::erm	SPP1 NRS5598 → NCIB3610
bdbCD	168 bdbCD::spc	Ref. 21
NRS5554	NCIB3610 bdbCD::spc	SPP1 NRS5139→ NCIB3610
NRS5553	NCIB3610 bdbCD::spc bdbA::erm	SPP1 NRS5139→ NRS5552
NRS5132	NCIB3610 bslA::cml amyE-Pspac-hy-gfpmut2 (spc) sacA::PbslA-bslA (kan)	SSP1 NRS1113→NRS2978
NRS5136	NCIB3610 bslA::cml amyE-Pspac-hy-gfpmut2 (spc) sacA::PbslA-bslA C178A C180A (kan)	SSP1 NRS5149→NRS5131
E. coli
MC1061	E. coli F'lacIQ lacZM15 Tn10 (tet)	E. coli Genetic Stock Centre
BL21 (DE3)	F– ompT hsdSB(rB–, mB–) gal dcm (DE3)	Ref. 46

^*Drug resistance cassettes are indicated as follows: cml, chloramphenicol resistance; kan, kanamycin resistance; mls, lincomycin and erythromycin resistance; and spc, spectinomycin resistance. BSGC represents the Bacillus genetic stock center.

^†The direction of strain construction is indicated with DNA or phage (SPP1) (→) recipient strain.

Genetic Separation of Roles in Biofilm Architecture and Hydrophobicity.

We next tested whether there was a biological consequence of restricting BslA disulfide bond formation in vivo. Deletion of bslA results in a flat, featureless, wetting biofilm; our analysis revealed that production of the BslA_C178A, BslA_C180A, BslA_C178A,C180A, and BslA_Δ172–181 variant proteins using the strains indicated could reinstate biofilm complexity of a bslA mutant to a level that was visually comparable to the wild-type strain (Fig. 1 C–I). We then assessed whether the biofilms formed were hydrophobic (8). As expected, the wild-type strain formed a nonwetting biofilm where the water droplet had a high contact angle (124.6 ± 2.9°), whereas the bslA mutant was wetting (33.6 ± 2.7°) (Table S2; refs. 8 and 12). Each of the single cysteine-to-alanine mutations, BslA_C178A (NRS5177) and BslA_C180A (NRS5178), displayed a nonwetting phenotype, although the contact angle calculated for the BslA_C178A (NRS5177) strain was consistently lower than that measured for the wild-type strain (P < 0.01; Student's t test) (Table S2). In sharp contrast, the architecturally complex biofilms formed in the presence of the monomeric BslA_C178A,C180A (NRS5179) or BslA_Δ172–181 (NRS2957) variants were wetting (Fig. 1 H and I), with contact angles that were indistinguishable from the bslA mutant strain (Table S2). Thus, monomeric BslA seems to be associated with architectural complexity of the biofilm, whereas surface hydrophobicity requires at least the dimeric form of the protein. We note that the single point mutations, BslA_C178A and BslA_C180A, yielded only dimeric protein in addition to the monomer (Fig. 1A), indicating that tetramer formation is not required. These results also reveal that the architectural complexity, which often makes a contribution to biological hydrophobicity (e.g., refs. 17 and 18), is not sufficient for hydrophobicity to manifest.

Table S2.

Contact angles of droplets placed on B. subtilis biofilms

Genotype†	Strain‡	Contact angle,§ °
Wild type	NCIB3610	124.6 ± 2.9
Wild type (lower surface)	NCIB3610	N.D.
bslA	NRS2097	33.6 ± 2.7
bslA + bslAWT	NRS2299*	106.9 ± 5.8
bslA + bslAAxC	NRS5177*	103.8 ± 2.6
bslA + bslACxA	NRS5178*	134.9 ± 2.2
bslA + bslAAxA	NRS5179*	34.1 ± 4.1
bslA + bslA∆Cterm	NRS2957*	29.3 ± 3.7
bslA + bslAWT : bslA + bslAAxA	NRS2299*:NRS5179* 1/1 ratio	116.2 ± 2.8
bslA + bslAWT : bslA + bslAAxA	NRS2299*:NRS5179*1/10 ratio	50.5 ± 1.3
bslA + bslAWT : bslA + bslAAxA	NRS2299*:NRS5179* 1/100 ratio	25.1 ± 0.9
bdbA	NRS5552	94.5 ± 3.8
bdbACD	NRS5553	54.8 ± 3.3
bdbCD	NRS5554	84.1 ± 3.2
bslA + bslAss-yweAWT	NRS5551*	34.8 ± 2.9
bslA + bslAss-yweACxC	NRS4834*	114.8 ± 1.1
bslA + bslAss-yweAAxC	NRS5515*	30.1 ± 1.2
bslA + bslAss-yweACxA	NRS5516*	34.7 ± 1.4
bslA + bslAss-yweAAxA	NRS5210*	21.7 ± 3.2
bslA#	NRS5131	30.5 ± 1.4
bslA + bslAWT#	NRS5132	119.6 ± 1.5
bslA + bslAAxA#	NRS5136	71.7 ± 2.7

^†Unless otherwise indicated, the upper surface of the biofilm was analyzed. With the exception of the values indicated with ^#, water was used as the test fluid. For those indicated with ^#, 1% (vol/vol) chlorhexidine gluconate was used.

^‡For the full strain details, see Table S1. All experiments were conducted by using 25 μM IPTG as required and indicated by *.

^§The average contact angle from a minimum of three independent samples is presented with the errors representing the SEM. N.D. represents samples where the contact angle was too low to measure.

Sharing BslA Molecules in the Biofilm.

Because hydrophobicity of the biofilms is at the macroscale, where BslA comprises a “common” or “public” good that can be shared by the population (19), we asked what minimum proportion of cells producing wild-type BslA was needed to achieve maximum nonwetting values. To determine this proportion, cells expressing either the wild-type bslA (NRS2299) or the bslA_C178A,C180A (NRS5179) coding regions were cocultured over a range of ratios, and the hydrophobicity of the resulting mature biofilm was measured (Fig. 1 J–L). When the strains were mixed in equal proportions, a nonwetting biofilm surface could be sustained (116.2 ± 2.8°). However, when 90% of the cells produced BslA_C178A,C180A, the biofilm hydrophobicity remained above that of the bslA mutant, but there was a step change in the wettability of the surface (50.5 ± 1.3°). Finally, wettability reached a value indistinguishable from that measured for the fully monomeric protein when 99% of the cells produced BslA_C178A,C180A (Table S2). These findings demonstrate that noncontributing bacteria can be tolerated during formation of the hydrophobic layer, as long as they represent <50% of the population.

Formation of the Hydrophobic Layer Depends on Thiol-Disulfide Oxidoreductases and Is Enhanced in an Oxic Environment.

In the B. subtilis extracytoplasmic space, disulfide bond formation is catalyzed by one of two thiol-disulfide oxidoreductases (TDORs) named BdbA and BdbD (20–22). We predicted that if BslA was actively oligomerized, then disruption of either bdbA or bdbD (or both in the case of functional redundancy) would produce a structured, but wetting, biofilm similar to the phenotype displayed by the monomeric BslA variants. To test this hypothesis, we constructed bdbA (NRS5552), bdbCD (NRS5554) (a bdbC bdbD operon deletion), and bdbA bdbCD (NRS5553) deletion strains and assessed biofilm formation and surface hydrophobicity (Fig. 2 A–C and Table S1). Consistent with our hypothesis, each of the strains formed biofilms that were morphologically indistinguishable from the parental strain NCIB3610 (Figs. 1C and 2 A–C), but measurement of surface wettability revealed that the double bdbA bdbCD mutant had a reduced contact angle of ∼55° compared with the wild-type strain (Table S2 and Fig. 2C). The single bdbA and bdbCD deletion strains produced surfaces with an intermediate level of hydrophobicity (Table S2 and Fig. 2 A and B).

Fig. 2.

The mechanism of disulfide bond formation. (A–C) Architecture and hydrophobicity of strains bdbA⁻ (NRS5552) (A), bdbCD⁻ (NRS5554) (B), and bdbACD⁻ (NRS5553) (C). Under each biofilm is an image of a water droplet on the upper surface of the biofilm; contact angle values are in Table S2. (D) Measurement of the oxygen concentration (μM) and redox potential (mV) as a function of the depth of the biofilm, with 0 mV being set at the biofilm surface (SI Materials and Methods for details). (E–G) Time course of water uptake by a mature biofilm visualized by using pigmented water: before treatment, 0 min (E); 2 min after exposure (F); and 15 min after exposure (G). In G, 5-mL colored water droplets demonstrate retention of upper biofilm hydrophobicity.

The wettability of the bdbA bdbCD mutant did not reach the level measured for the bslA deletion strain (Table S2). Therefore, unless there is a redundant enzymatic mechanism, we propose that a combination of catalytic and spontaneous disulfide bond formation drives dimerization of BslA and leads to the formation of the hydrophobic coat. Consistent with this hypothesis, measurement of the oxygen concentration through the biofilm by using a microsensor demonstrated a steep decrease in the available oxygen, with no oxygen measured after 50–60 μm, as measured from the air-exposed surface, rendering the lower biofilm region anoxic (Fig. 2D). Furthermore, microelectrode-based assessment of the change in redox potential through the biofilm indicated that the upper region of the biofilm community was more oxidizing than the base (Fig. 2D). The lack of electron acceptors at the base of the biofilm suggests that BslA may be in a monomeric form in this region. Given the observation that surface hydrophobicity is associated with the ability to form dimeric protein (Fig. 1 C–I and Table S2), a predicted prevalence of monomeric protein at the base of the biofilm raises the possibility that this surface will be wetting. Two experimental findings supported this hypothesis: First, when pigmented water was flooded onto a Petri dish on which a biofilm had been grown, the dye was observed to move rapidly under the biofilm toward the center of the community (Fig. 2 E–G), indicating that it was not repelled by a hydrophobic layer. Second, when the mature wild-type biofilm was turned over, such that the base was facing up, an entirely wetting surface was revealed (Table S2). Together, these findings solve the conundrum (12) of how bacteria in the biofilm access nutrients, despite being surrounded by a layer of BslA.

BslA Localization Depends on Disulfide Bond Formation.

We next addressed the mechanism by which blocking oligomerization of BslA impaired the formation of a nonwetting biofilm. It is not a consequence of protein misfolding or disruption of protein production in vivo, because CD spectroscopy demonstrated similar secondary structures in the variant proteins (Fig. S2A), and immunoblot analysis revealed comparable levels of the BslA cysteine mutants to the wild-type protein (Fig. 1A). Therefore, we considered two nonmutually exclusive hypotheses: (i) Mutation or deletion of the C-terminal cysteine residues impairs the innate ability of BslA to form a stable elastic film that depends on lateral interactions between the monomers (12), and consequentially the ability to render the biofilm nonwetting; and (ii) BslA_C178A,C180A cannot form a dense layer over the surface of the biofilm as observed for the wild-type protein (12). The ability of recombinant BslA_C178A, BslA_C180A, BslA_C178A,C180A, and BslA_Δ172–181 to form stable elastic protein films in vitro was assessed by using pendant drop analysis, coupled with quantification of the wrinkle relaxation speed (12). Each of the variant proteins formed a stable protein film at the oil–water interface with no significant differences compared with wild-type BslA (Fig. S2B). Furthermore, transmission electron microscopy (TEM) indicated that the ability of the variant proteins to form an ordered 2D lattice at an interface was not impeded (Fig. S2 C–E).

Next, in situ localization of BslA was assessed by using immunofluorescence staining of biofilm cross-sections coupled with confocal microscopy (12). The strains tested were the wild type, the bslA mutant, and the bslA mutant complemented with either the wild-type or bslA_C178A,C180A coding region. In each case, the strains were modified to express the gfp coding region to allow detection of the cells by confocal microscopy (Table S1). As has been observed (12), both the wild-type strain (Fig. S3A) and the bslA mutant complemented with the wild-type bslA allele, showed BslA-associated fluorescence at both the air–cell and cell–agar interfaces, consistent with formation of the BslA integument (Fig. 3A and Fig. S3C). Specificity of the immunofluorescence staining was confirmed by analysis of the bslA mutant (Fig. S3B). In contrast, when the bslA mutant was complemented with the BslA_C178A,C180A variant (NRS5136), BslA-linked fluorescence was not detected in high abundance at the cell–air interface and was only prevalent at the agar–cell junction (Fig. 3B and Fig. S3D). These findings indicate an inability of BslA_C178A,C180A either to migrate to or accumulate at the air interface, which is consistent with the wetting phenotype of the biofilm formed when this variant of BslA is produced.

Fig. S3.

In situ analysis of BslA localization in the biofilm. Confocal scanning laser microscopy images of cross-sections through biofilms formed by wild-type cells (NRS1473) (A), the bslA mutant strain (NRS3812) (B), the bslA mutant complemented with either the wild-type variant of bslA (NRS5132) (C), or the gene encoding the BslA_AxA variant (NRS5136) (D) are shown. For the merged images, the fluorescence from the GFP within the cells is shown in green, and the fluorescence associated with DyLight594, representing immunolabeled BslA staining, is shown in magenta. (Scale bar: 50 µm.)

Fig. 3.

In situ analysis of BslA localization in the biofilm. Confocal scanning laser microscopy images of biofilm cross-sections through the bslA mutant complemented with either the wild-type variant of bslA (NRS5132) (A) or the gene encoding the BslA_{C178A C180A} variant (NRS5136) (B) strains are shown. Fluorescence from the GFP is shown in green, and the fluorescence associated with DyLight594, representing immunolabeled BslA staining, is in magenta. (Scale bars: 50 µm.) The single-channel data and wild-type and bslA controls are in Fig. S3.

Synthetic Dimerization of the BslA Paralogue.

The strict requirement for dimeric BslA to mediate biofilm hydrophobicity led us to hypothesize that if the monomeric paralogue of BslA, YweA, could be engineered to dimerize in vivo, it may become capable of reinstating the nonwetting phenotype to the bslA mutant. This outcome is not the case when the wild-type yweA coding region is expressed from a heterologous location in the bslA mutant (14). To test this prediction, we synthesized a chimeric YweA allele (Fig. S4A), fusing the C-terminal 11 amino acids of BslA (containing C178 and C180) to the C terminus of YweA (hereafter YweA_BslA171-181). We additionally generated chimeric constructs, where each of the cysteine residues in the BslA C-terminal region was individually, and in combination, mutated to alanine (Table S1). The wild-type YweA, chimeric YweA_BslA171-181, and YweA_{BslA171-181 C178A C180A} recombinant proteins were purified from Escherichia coli, and the secondary structure was assessed by CD spectroscopy. This analysis revealed that the C-terminal extension did not significantly affect folding of the protein compared with the parental YweA protein (Fig. S5A). The oligomerization state of the YweA_BslA171-181 chimeric protein was assessed in vitro by SDS/PAGE (Fig. S4 B and C) and size-exclusion chromatography (SEC) (Fig. S4D), which showed that the chimeric YweA_BslA171-181 protein formed dimers that were reduced to the monomeric state by the addition of β-mercaptoethanol. Bands with the apparent molecular weight expected for dimers were also formed when either one of the two cysteine residues remained (YweA_{BslA171-181 C178A} and YweA_{BslA171-181 C180A}) (Fig. S4B), but only monomer was observed for the double cysteine-to-alanine chimeric protein (YweA_{BslA171-181 C178A C180A}) (Fig. S4B). Unlike BslA, we did not detect formation of tetramers for the YweA_BslA171-181 chimeric protein. Notably, fusion of the BslA C-terminal 11 amino acids to YweA did not alter the stability or organization of the protein film formed in vitro. Pendant drop analysis, coupled with quantification of wrinkle relaxation speed, revealed that YweA, YweA_BslA171-181, and YweA_{BslA171-181 C178A C180A} each had an average relaxation time of ∼25 s (Fig. S5B), substantially less stable than the BslA elastic film (>600 s; Fig. S5B). This result means that the proteins can form elastic films, but they are unstable under compression. Consistent with the film stability, TEM showed that each protein was able to form an organized lattice on a surface (Fig. S5 C–E). Together, these findings indicate that we can generate YweA oligomers in vitro by the addition of the BslA C-terminal 11 amino acids, but although they form an ordered 2D lattice, the chimeric proteins retain the fast film-relaxing properties of YweA (14).

Fig. S4.

In vitro and in vivo analysis of recombinant YweA and chimeric YweA protein higher-order forms. (A) Schematic of the YweA chimeric forms: SS represents the BslA signal sequence used for export, and CxC indicates the C-terminal 11 amino acids from BslA. (B and C) SDS/PAGE analysis of 30 μg of YweA, YweA_BslA171-181, YweA_{BslA171-181 C178A}, YweA_{BslA171-181 C180A}, and YweA_{BslA171-181 C178A C180A} recombinant protein in the absence (B) and presence (C) of reducing agent. (D) SEC analysis of recombinant proteins using a Superdex 75 10/300GL column in an unreduced state. (E) Western blot analysis of YweA in a reduced state detected from biofilm protein extracts.

Fig. S5.

Transplantation of the CxC motif to the YweA C terminus does not restore in vitro BslA behavior. (A) Solution state CD spectra of WT YweA (black), YweA_BslA171-181 (red), and YweA_{BslA171-181 C178A C180A} (blue) indicate that all three variants have a similar secondary structure. (B) Wrinkle relaxation of YweA (black), YweA_BslA171-181 (red), and YweA_{BslA171-181 C178A C180A} (blue). Wrinkles were formed in the protein film by removing fluid from the droplet, and relaxation of the wrinkles was followed over time. The error bars represent the SD in normalized greyscale over at least five independent wrinkles. For all three variants, wrinkle relaxation was very rapid. (C–E) TEM images of YweA (C), YweA_BslA171-181 (D), and YweA_{BslA171-181 C178A C180A} (E) stained with uranyl acetate indicate that all three variants are capable of forming the ordered 2D lattice observed previously for WT BslA (11, 12) and YweA (44).

Oligomeric YweA Yields a Hydrophobic Biofilm.

To assess the impact of engineering YweA to form intermolecular disulfide bonds in vivo, we constructed a suite of plasmids designed to produce the YweA chimeric proteins in B. subtilis. The constructs were generated such that secretion through the Sec-system was directed by the BslA signal sequence (the variants are hereafter referred to as YweA_BslA171-181, YweA_{BslA171-181 C178A,} YweA_{BslA171-181 C180A}, and YweA_{BslA171-181 C178A C180A}) (Fig. S4A). The plasmids carrying the required coding regions were introduced into the bslA mutant at the ectopic amyE locus (Table S1). Western blot analysis of proteins extracted from the biofilm, by using an anti-YweA antibody, revealed that only in the presence of both cysteine residues in the appended BslA C terminus could significant levels of disulfide bond-dependent oligomeric chimeric YweA protein be detected (Fig. 4A). As anticipated, the addition of a reducing agent during the extraction process rendered the protein fully monomeric (Fig. S4E). The ability of each of the YweA chimeric variants to reinstate biofilm structure and hydrophobicity to the bslA mutant was tested as before. As previously observed, induction of wild-type yweA expression did not alter the biofilm morphology or wetting phenotype, compared with the parental bslA mutant (Fig. 4B) (14). In sharp contrast, appending the BslA C-terminal 11 amino acids to YweA allowed the chimeric YweA_BslA171-181 protein to return hydrophobicity to the bslA mutant strain (114.8 ± 1.1°) (Fig. 4B and Table S2). It should, however, be noted that the biofilm structure retained more architectural similarities with the bslA mutant, rather than the wild-type NCIB3610 strain (Fig. 4B). Expression of YweA_BslA171-181 thus results in an unstructured, yet nonwetting, phenotype. The slightly lower contact angle measured (114.8 ± 1.1° vs. 124.6 ± 2.9° for the structured, wild-type biofilms) may indicate that biofilm architecture makes some contribution to overall hydrophobicity, as has been suggested elsewhere (18). In contrast, when the C-terminal extension was mutated such that one (NRS5515 and NRS5516) or both (NRS5210) cysteine residues were replaced with alanine, yielding monomeric BslA in vivo, the colonies formed retained both the wetting and unstructured phenotypes exhibited by the bslA mutant (compare Fig. 1D with Fig. 4B). Thus, dimeric chimeric YweA containing the two cysteine residues is able to reinstate biofilm hydrophobicity, whereas monomeric chimeric YweA is unable to reinstate biofilm architectural complexity.

Fig. 4.

Forcing dimerization of YweA can rehabilitate biofilm hydrophobicity in a bslA mutant. (A) Western blot analysis of YweA in a native state detected from biofilm protein extracts (SI Materials and Methods). Strains used were 3610 (WT; NCIB3610), yweA⁻ (NRS2405), and the bslA mutant engineered to produce the following YweA variants: YweA (NRS5551), YweA_BslA171-181 (NRS4834), YweA_{BslA171-181 C178A} (NRS5515), YweA_{BslA171-181 C180A} (NRS5516), and YweA_{BslA171-181 C178A C180A} (NRS5210). (B) Biofilms using strains detailed in A: Under each biofilm is an image of a water droplet on the upper surface of the biofilm: values of the contact angle are in Table S2 and strains are as in A.

Hydrophobicity and Architecture Play a Role in Resistance to Chemical Attack.

The ability to genetically separate biofilm structure from hydrophobicity allowed us to assess whether protection from external insult is conferred to the bacteria by blocking access of the chemical and/or mediated by the architectural complexity. B. subtilis biofilms are nonwetting when challenged with selected commercial biocides (6), with chlorhexidine gluconate being the reactive agent of a number of such biocides. Analysis revealed that 1% (vol/vol) chlorhexidine gluconate is nonwetting on “wild-type” biofilms (Fig. 5A) and, in contrast, is wetting on both the bslA mutant and the variant carrying the monomeric BslA_C178A,C180A (Fig. 5A and Table S2), providing an ideal test agent to test the question posed above. The number of surviving cells after treatment for 5 min with 1% (vol/vol) chlorhexidine gluconate was calculated relative to a comparable sample exposed to saline solution. We noted a clear role for BslA in mediating cell survival: In the absence of bslA, exposure to 1% (vol/vol) chlorhexidine gluconate resulted in ∼38-fold fewer surviving cells compared with when BslA was produced (Fig. 5B). In contrast, when BslA_C178A,C180A was present, resulting in a structured, but wetting, biofilm, limited cell survival was measured. Cell survival was approximately fivefold lower than that measured for the wild-type strain (Fig. 5B). Because BslA is known to have a role in sporulation, specifically during biofilm formation (19), we calculated the level of heat-resistant spores in each biofilm at the time point used to test chemical resistance. As established previously, the level of sporulation was low in the bslA mutant (Fig. 5C); however, the number of spores formed in the presence of BslA_C178A,C180A was the same as when wild-type BslA was produced (Fig. 5C). In toto, these data demonstrate that the protection received by the cells in the biofilm results from a combination of the hydrophobicity and structure conferred by BslA, a phenomenon only revealed by the ability to genetically separate the two functional contributions.

Fig. 5.

Cell survival after exposure to chlorhexidine gluconate. (A) An image of a 1% (vol/vol) chlorhexidine gluconate droplet on the upper surface of the biofilm for strain WT (NRS5132), bslA mutant (NRS5131), and the bslA mutant complemented with BslA_{C178A C180A} +AxA (NRS5136); the values of the contact angle are in Table S2. (B) The strains described above were exposed to 1% (vol/vol) chlorhexidine gluconate, and the percentage survival was calculated. (C) Percentage sporulation was calculated for the strains detailed in A. The error bars represent the SD from the mean.

Discussion

The mechanism revealed here for controlling protein function through oxidation allows us to classify BslA as a bifunctional protein with genetically separable roles in biofilm formation, mediated by cysteine residues in the C terminus. BslA is a key component of the B. subtilis biofilm and plays a role in both biofilm architecture and hydrophobicity (8, 12, 23). Dimerization of BslA directly facilitates the development of the nonwetting layer and relies on a cysteine motif (CxC) at the C terminus. Monomeric variants of BslA yield biofilms that are architecturally complex, but lack hydrophobicity, whereas, conversely, transplantation of the BslA C terminus onto the normally monomeric nonfunctional paralogue YweA allows the chimeric protein to dimerize and consequentially rehabilitate hydrophobicity, but not architecture, to a bslA mutant. These findings are consistent with the accumulation of BslA at the air–biofilm interface only when it can dimerize, as observed by high-resolution in situ immunofluorescence microscopy (Fig. 3). The BslA layer is thus shared by the entire community and can form over and shield nonproducing bacteria (Fig. 1 J–L). By virtue of being able to separate the role of BslA in biofilm architecture from hydrophobicity, we have shown that both architectural complexity and the hydrophobic layer contribute to protecting the residents from biocides, thus providing an evolutionary advantage to the resident cells. Finally, we have demonstrated that an effective hydrophobic barrier can be generated by a subpopulation of the biofilm, and this finding raises the question of why the whole population has evolved to produce BslA (12) when other biofilm matrix molecules are produced in a bimodal manner (24).

A model can be postulated that explains formation of a BslA layer that is more than one molecule (or dimer) deep at the air interface (Fig. 6). It is known that BslA dimers are orientated longitudinally (tail-to-tail), with only one cap of each dimer able to interact with the air interface at a time (11). The orientation of tetrameric forms of the protein is unclear, however; because dimeric protein is sufficient to give rise to the observed effects, it is not considered further here. Based on the premise that the hydrophobic cap of one dimer can serve as a hydrophobic interface for another dimer, it leads to a scenario where the integument of the biofilm could comprise the dimeric proteins layered against each other (Fig. 6). Furthermore, because monomeric and dimeric BslA are able to coexist in vitro in a protein film, the mature in vivo hydrophobic layer has the potential to be generated from a mixture of monomeric and dimeric protein. We have confirmed that stable in vitro film formation is not needed to confer hydrophobicity; the chimeric YweA_{BslA171–181} variant is able to render the community hydrophobic, but forms a protein film that relaxes quickly under compression (Fig. 4B and Fig. S5B). These findings are consistent with a recent analysis of BslA orthologs, where a Bacillus pumilus variant lacked stability in the elastic film, but could substitute for the B. subtilis bslA gene in vivo (14). Together, these data indicate that stability in the lateral interactions between the BslA molecules is independent of the ability to confer hydrophobicity to the community, but may determine architectural complexity. However, details of the lateral interaction sites between the protein molecules are currently unknown.

Fig. 6.

Model of BslA function. (i) Schematic of a biofilm cross-section depicting the oxygen and redox gradient through the depth of the structure. The blue layer at the air interface represents the BslA hydrophobic coat with a water droplet on top (red circle), and the hatched lines at the base represent BslA in a form that allows water and nutrient uptake into the community. The bacteria are green ovals, and the agar surface is the gray zone at the base of the biofilm. (ii) Oligomerization of BslA is mediated by thiol-oxidoreductases that reside in the membrane and extracytoplasmic space. The electrons (e⁻) released by oxidation upon formation of the disulfide bond flow into the respiratory chain. BslA is shown as a blue oval in the biofilm matrix (gray). For simplicity, only one disulfide bond has been depicted (S–S). The reduced form of the protein is represented by the (−SH) annotation. (iii) Oligomerization of BslA is also likely to occur using molecular oxygen as the electron acceptor. (iv) Depicted is a model for how the BslA coat might present in the biofilm as a mixture of oligomers. The ability of BslA to present in a cap-in and -out configuration is represented, where the cap-out form is adopted by the molecules at a hydrophobic interface.

BslA is therefore multifunctional across three different axes: (i) Dimerization (and/or tetramerization) contributes to hydrophobicity, whereas monomeric protein is sufficient for biofilm complex architecture; (ii) the cap-out form of the protein renders a surface layer hydrophobic, whereas we can infer that a cap-in form is present in the wetting protein layer at the base of the biofilm where the protein is exposed to water (11, 13); and (iii) stable lateral interactions between BslA molecules, which can be measured in vitro, appear to be required for architectural complexity, whereas unstable lateral interactions (reflected in unstable film formation in vitro) nonetheless are sufficient to give rise to hydrophobicity if dimeric protein is present. We have previously elucidated a link between the behavior of the cap region and the strength of lateral interactions (13), such that these two functionalities appear to influence each other. Conversely, the behavior of the cap region of BslA is functionally independent from the protein oligomerization state, because both monomeric and dimeric protein are able to flip between cap-in and -out conformations. Thus, the bottom of the biofilm is wetting not just because the protein is monomeric, but because the cap region is also exposed to water; likewise, the top of the biofilm is hydrophobic, not just because BslA forms dimers and tetramers, but because the cap region is also exposed to air. The mechanism(s) by which monomeric BslA gives rise to an architecturally complex biofilm, and the reasons why dimeric protein (at least) is required to give rise to a hydrophobic coat, remain to be elucidated. We have previously observed a correlation between architectural complexity of the biofilm and the ability of BslA variants to form a stable elastic film (14); these previous results are supported by our current findings that chimeric YweA, which forms an unstable film in vitro, can reinstate biofilm hydrophobicity, but not complex architecture. We speculate that the stable elastic film formed by monomeric BslA facilitates an association with another component(s) of the biofilm matrix—for example, the exopolysaccharide. Moreover, consistent with this proposal, an association between different molecular components of the B. subtilis biofilm matrix can be presumed, because loss of any individual element leads to impaired structure (19).

Bifunctionality Through Dimerization.

Cysteine residues, possessing a thiol group (15), have specialized roles in a wide range of cellular processes and control protein folding and stability, multimerization, and function through disulfide bond formation (25). It is clear from our secondary structural analysis (Fig. S2) that the role for disulfide bond formation in modulating BslA activity is not linked with either protein stability or folding, but is instead associated with imparting new function through multimerization. This function is in contrast to the fungal hydrophobins, where disulfide bonds stabilize the structure of the protein (26), and is more analogous to the mechanism used to control activity of the von Willebrand factor during blood clotting (25). Previous bioinformatic analyses revealed that Firmicutes, including B. subtilis, limit the number of proteins that contain cysteine residues (27). Consistent with this conclusion, there are no essential components that require disulfide bond formation in the B. subtilis biofilm, because deletion of the known extracytoplasmic thiol oxidoreductases, bdbA and bdbD, does not lead to pleiotropic defects. It is only when the integrity of hydrophobicity is assessed that differences in the surface wettability are uncovered (Table S2). It has been proposed that in Firmicutes, proteins that contain disulfide bonds have roles in “niche” functions that are not linked with essential cellular processes—for example, genetic competence that allows the uptake of exogenous DNA (21, 28) and production of the S-linked glycopeptide sublancin that has antimicrobial properties (23, 29, 30). Arguably, biofilm hydrophobicity would fit within this category, and, consistent with genetic competence and sublancin production, successful formation of the hydrophobic layer imparts a fitness advantage by, in this case, excluding chemicals from entering the biofilm.

Heterogeneity in Electron Acceptor Availability Drives BslA Function.

In addition to catalysis-driven disulfide bond formation by extracytoplasmic thiol-oxidoreductases, spontaneous disulfide bond formation using molecular oxygen (or via a redox active molecule) as the direct electron acceptor enhances BslA dimerization at the air–biofilm surface interface. This process is driven in the top layer of the biofilm by an oxidative, oxygen-rich zone that is conducive to disulfide bond formation (Fig. 6) (31). It is well established that oxygen gradients stratify natural, mixed-species biofilms by driving distribution of microorganisms with different respiratory requirements (32–34). Furthermore, the availability of electron acceptors, including oxygen, can drive structuring of biofilm morphology, as has been shown for Pseudomonas aeruginosa (35, 36). Therefore, these findings expand the role that oxygen gradients can play in biofilm formation by highlighting an ability to modulate protein function. The ability to respond to localized oxygen heterogeneity in this manner provides an efficient mechanism for bacteria to maximize resource utilization by generating bifunctionality through redox sensitivity. Furthermore, BslA dimerization in response to the oxygen gradient solves the conundrum of how B. subtilis obtains nutrients when it is surrounded by a layer of BslA (12) [i.e., the anoxic base of the biofilm is hydrophilic despite the abundance of BslA (Fig. 6)]. We predict that bifunctionality of matrix components will emerge as a common theme across the species. Consistent with this proposal, the Vibrio cholerae biofilm matrix protein, RmbA, is proteolytically cleaved to a form that promotes recruitment of cells to the biofilm in an exopolysaccharide-independent manner; effectively, the proteolytic event changes the function of RmbA (37). Additionally, flagella synthesized by E. coli are used, not only for motility, but also for imparting structural rigidity to the community by entwining the bacterial cells (38). Control of bifunctionality for each of these examples is distinct and raises the question of how many different mechanisms have evolved to maximize use of a limited number of components.

Materials and Methods

Details of all methods used are provided in full in SI Materials and Methods.

Growth Conditions.

The B. subtilis strains used and constructed in this study are detailed in Table S1. The full details of growth conditions are provided in SI Materials and Methods. Biofilm colonies were grown on MSgg medium (5) solidified with 1.5% Select Agar (Invitrogen) at 30 °C for 48 h.

Strain Construction.

All strains, plasmids, and primers used are presented in Tables S1, S3, and S4 and were constructed by using standard techniques. See SI Materials and Methods for full details.

Table S3.

Plasmids used in this study

Plasmid	Description*	Source†
pDR111	B. subtilis amyE integration vector for IPTG-induced expression	Ref. 47
pGEX-6P-1	Vector for production of GST-fused proteins	GE Healthcare
pQE70	Cloning vector for His-tag fusions	Qiagen
pET15b-TEV	Vector for production of His tag-fused proteins	Lab sources
pNW1420	pET15bTEV-yweA31–155	This work
pSac-Kan	B. subtilis sacA integration vector	Ref. 48
pNW1500	pDR111-bslAC178A	This work
pNW1501	pDR111-bslAC180A	This work
pNW1502	pDR111-bslAC178A C180A	This work
pNW1128	pGEX-6P-1-TEV-bslA42–181	Ref. 12
pNW1503	pGEX-6P-1-TEV-bslA42–181 C178A	This work
pNW1504	pGEX-6P-1-TEV-bslA42–181 C180A	This work
pNW1505	pGEX-6P-1-TEV-bslA42–181 C178A C180A	This work
pNW1506	pET15b-TEV-yweA-bslA171–181	This work
pNW1507	pDR111-bslAss-yweA-bslA171–181 C178A C180A	This work
pNW1508	pET15b-TEV-yweA-bslA171–181 C178A C180A	This work
pNW1510	pGEX-6P-1-TEV-bslA42–171 (Δ172–181)	This work
pNW1700	pET15b-TEV-yweA-bslA171–181 C178A	This work
pNW1701	pET15b-TEV-yweA-bslA171–181 C180A	This work
pNW1702	pDR111-bslAss-yweA-bslA171–181 C178A	This work
pNW1703	pDR111-bslAss-yweA-bslA171–181 C180A	This work
pNW518	pSac-Kan-PbslA-bslA	This work (coding region amplified with primers NSW646 and NSW645 cloned into pSac-Kan)
pNW1231	pSac-Kan-PbslA-bslAC178A C180A	This work (C178A and C180A mutations introduced into pNW518 with primers NSW1124 and NSW1125)
pNW1079	pDR111- bslAss- yweA-bslA171–181	This work (coding region from pNW1075 cloned into pDR111 using HinDIII and SphI)
pNW611	pQE70-bslA signal sequence (hereafter bslAss)	Ref. 19
pNW1075	pQE70-bslAss-yweA-bslA171–181	This work (synthetic gene amplified with primers NSW2026 and NSW645 and cloned into pNW611)
pNW512	pDR111-bslA	Ref. 19
pNW621	pDR111-bslAΔ172–181-	This work (coding region amplified with primers NSW626 and NSW838)
pNW1718	pQE70- bslAss-yweA	This work
pNW1719	pDR111- bslAss- yweA	This work

^*The genotype of the plasmid is described.

^†Relevant information for the construction of the plasmid is provided, along with the references if previously published.

Table S4.

Oligonucleotide primers used in this study

Primer	Sequence 5′–3′†	Use‡
NSW12	CGATTCAAAACCTCTTTACTG	amyE locus for assessing double crossovers
NSW13	GCTTAAGCCCGAGTC	amyE locus for assessing double crossovers
NSW1853	GTACCATATGCAGTCTGCATCAATCGAG	yweA31–155 cloning into pET15b-TEV (leaves GHM at N-term end of YweA)
NSW1854	GATCCTCGAGTTATTAACGGGGATCAATCAC	yweA31–155 cloning into pET15b-TEV (leaves GHM at N-term end of YweA)
NSW1900	TCCTCCGACTCAGCCTgcaGGTTGCAACTAAGCAT	SDM on pNW512 to convert C178 to Ala
NSW1901	ATGCTTAGTTGCAACCtgcAGGCTGAGTCGGAGGA	SDM on pNW512 to convert C178 to Ala
NSW1902	GACTCAGCCTTGCGGTgcaAACTAAGCATGCAAGC	SDM on pNW512 to convert C180 to Ala
NSW1903	GCTTGCATGCTTAGTTtgcACCGCAAGGCTGAGTC	SDM on pNW512 to convert C180 to Ala
NSW1904	TCCGACTCAGCCTgcaGGTgcaAACTAAGCATGCA	SDM on pNW512/pNW1079 to convert C178 and C180 to Ala

^†Restriction sites are underlined, and sites for mutations are highlighted in lowercase bold.

^‡Relevant information for the use of the primers is provided. SDM, site-directed mutagenesis.

SI Materials and Methods

General Growth Conditions and Strain Construction.

The B. subtilis and E. coli strains used and constructed in this study are detailed in Table S1. E. coli strain MC1061 was used for the construction and maintenance of plasmids. B. subtilis 168 derivatives were obtained by transformation of competent cells with plasmids using standard protocols (39). SPP1 phage transductions were used to introduce DNA into B. subtilis strain NCIB3610 (40). Both E. coli and B. subtilis strains were routinely grown in Lysogeny Broth (LB) medium (10 g of NaCl, 5 g of yeast extract, and 10 g of tryptone per liter) at 37 °C for 16 h. For complex colony formation, B. subtilis strains were grown on MSgg medium (5 mM potassium phosphate and 100 mM Mops at pH 7.0 supplemented with 2 mM MgCl₂, 700 μM CaCl₂, 50 μM MnCl₂, 50 μM FeCl₃, 1 μM ZnCl₂, 2 μM thiamine, 0.5% glycerol, and 0.5% glutamate) (5) solidified with 1.5% Select Agar (Invitrogen) at 30 °C for 48 h (5) and imaged as described (40). Ectopic gene expression was induced by medium supplementation with 25 µM isopropyl β-d-1-thiogalactopyranoside (IPTG) as indicated. When appropriate, antibiotics were used at the following concentrations: ampicillin 100 μg⋅mL⁻¹, chloramphenicol 5 μg⋅mL⁻¹, erythromycin 1 μg⋅mL⁻¹ with lincomycin 25 μg⋅mL⁻¹, kanamycin 25 μg⋅mL⁻¹, and spectinomycin 100 μg⋅mL⁻¹. ΔbdbA, ΔbdbCD, and ΔbdbACD mutant strains were constructed as described in Table S1 by using standard methodologies.

Plasmid Construction and Site-Directed Mutagenesis.

All strains, plasmids, and primers used in this study are presented in Tables S1, S3, and S4 and were constructed by using standard methods. The plasmids for BslA_42–181 overproduction were obtained by site-directed mutagenesis using the plasmid pNW1128 as template, which is a pGEX-6P-1 derivative used previously to overexpress BslA_42–181 (12). Primers for the codon substitutions are included in Table S4, and mutagenesis was achieved following the Stratagene QuikChange kit recommendations. The plasmids for overexpression yweA or the fusion yweA-BslA_171–181 were obtained by using standard techniques with either NCIB3610 or a synthetic DNA construct as the template DNA during PCR. The sequence of the synthetic construct generated by Genescript was as follows: 5′-agatct CGA TCT gca tca atc gag gca aaa aca gtt aac agc acg aaa gag tgg acc att tct gat att gaa gtg aca tat aaa cca aat gcg gtg ctt tct ctt gga gcg gta gaa ttt caa ttt cct gac ggg ttt cat gct acg aca aga gat tca gtg aat gga aga aca ctg aaa gaa aca cag att tta aac gat gga aaa aca gtc aga ctc ccg ctt acg ctt gat ttg tta ggc gca tcc gaa ttt gac ctt gtc atg gtg cgt aaa act ctt cct cgc gca ggc act tac acg att aaa ggc gat gta gta aac ggt ttg gga atc ggc agt ttt tat gct gaa acg cag ctg gtg att gat ccc cgt agc act cct ccg act cag cct tgc ggt tgc aac taa GCA TGC-3′. The sequence in capital letters represents restriction sites, the sequence in bold represents the C-terminal bslA coding region, and, finally, the sequence in lowercase represents the mature region of the yweA gene sequence.

Biofilm Hydrophobicity Contact Angle Measurements.

Biofilm hydrophobicity was evaluated by measuring the contact angle of a 5-µL droplet of water or 1% (vol/vol) chlorhexidine gluconate placed on the upper or lower surface of the biofilm that had been grown for 48 h at 30 °C. Measurements were obtained by using a ThetaLite TL100 optical tensiometer (Biolin Scientific) and analyzed with OneAttension. The water droplet was allowed to equilibrate for 5 min before imaging and measurement. Contact angles are present as the average of at least three independent experiments and the SEM associated with these values (Table S1).

Protein Purification.

BslA_41–181 protein and its derivatives were overexpressed and purified as described (12). Briefly, the pGEX-1–6P derivative plasmids were introduced into E. coli BL21 (DE3). After growth and overexpression, the protein was purified in Hepes buffer. To achieve this purification, the E. coli cells were lysed in an Emulsiflex cell disruptor, and the solubilized protein extracts, cleared of cell debris, were incubated with Glutathione Sepharose 4B (GE Healthcare), allowing the fused protein to bind to the GST binding beads. After incubation, the beads containing the BslA-fusion were recovered by using a gravity flow column (Bio-Rad) and suspended into new buffer containing DTT and TEV–His-tagged protease. TEV removed the GST tag from BslA protein, which remained soluble in the purification buffer. The protease and unbound GST were then separated from BslA by incubating the mixture with Ni-nitrilotriacetic acid (Ni-NTA) agarose (Qiagen) and glutathione beads, followed by a new passage in a gravity flow column. The flow-through recovered contained the purified protein, which was then concentrated by using VivaSpin concentrator. Protein quality was confirmed by SEC analysis.

To purify YweA_31–155 and the chimeric proteins YweA-BslA_171–181 and its derivatives YweA-BslA_{171–181 C178A}, YweA-BslA_{171–181 C180A}, and YweA-BslA_{171–181 C178A C180A}, the required coding regions were introduced into pET-15b-TEV vector (Tables S3 and S4). The pET-15b-TEV derivative plasmids were introduced into E. coli BL21 (DE3). After growth and overexpression, E. coli cells were suspended in Hepes buffer (300 mM NaCl and 50 mM Hepes, pH 8, with the addition of 20 mM imidazole) containing protease inhibitors (Roche cOmplete EDTA-free) and lysed by using the Emulsiflex C3. The solubilized protein extracts, cleared of cell debris, were incubated with Ni-NTA agarose (Qiagen) for 4 h, allowing the fused protein to bind to the Ni-NTA binding beads. After incubation, the beads containing the YweA-fusion proteins were recovered by using a gravity flow column (Bio-Rad) and suspended into new buffer (300 mM NaCl and 50 mM Hepes, pH 8.0, with the addition of 250 mM imidazole) on column for a half-hour to promote the protein release. Proteins were recovered by collection of the column flow-through, which was incubated overnight in buffer 300 mM NaCl and 50 mM Hepes (pH 8.0), with the addition of DTT and TEV–His-tagged protease. VivaSpin concentrators were used to remove the imidazole in solution by replacement with the purification buffer. The protein recovered from the concentrators was incubated overnight with Ni-NTA beads to bind to the His-tag fragments released, followed by a new passage in a gravity flow column. The flow-through recovered contained the purified protein, which was then concentrated by using VivaSpin concentrators. Protein quality was confirmed by SEC analysis.

SEC.

To evaluate the presence or absence of dimers in the purified proteins, 500 ng of purified protein was analyzed by SEC using a Superdex 75 10/300GL column with a low rate of 0.5 mL/min in the original purification buffer. Protein samples were also analyzed by using 14% SDS/PAGE, with and without the addition of a reducing agent (β-mercaptoethanol) as the loading dye and stained with Instant Blue before photography.

CD Analysis.

CD analysis was performed by using a Jasco J-810 spectropolarimeter. Samples were measured in a 0.1-cm quartz cuvette at 0.15 mg/mL protein concentrations in 25 mM phosphate buffer (pH 7). All samples were measured by using a 50 nm/s scan rate, 0.1-nm data pitch, and a digital integration time of 50 nm/s. Twenty individual spectra were measured and averaged per sample.

TEM.

A 5-μL droplet of 0.025 mg/mL protein solution (in 25 mM phosphate buffer, pH 7) was pipetted onto a carbon-coated copper grid (TAAB Laboratories Equipment Ltd.) and left for 5 min before being wicked away from the side by using filter paper. Subsequently, a 5-μL droplet of 2% uranyl acetate was pipetted onto the grid. This droplet was also left for 5 min before being wicked away from the side. Stained grids were then imaged by using a Philips/FEI CM120 BioTwin transmission electron microscope.

Wrinkle Relaxation.

The 0.2 mg/mL protein samples (in 25 mM phosphate buffer, pH 7) were loaded into a glass syringe with a needle diameter of 1.83 mm. A 40-μL droplet of protein solution was expelled into glycerol trioctanoate oil and allowed to equilibrate for 20 min at room temperature. Subsequently, the droplet was compressed by retraction of 7 μL from the droplet, resulting in the formation of wrinkles in the protein film. Images of the droplet were taken from the moment of compression by using a CCD camera. Images were acquired at a rate of 30 frames per second (fast relaxers; all YweA variants) or 2 frames per second (slow relaxers; all BslA variants) for up to 10 min. Relaxation of the wrinkles was analyzed in ImageJ. A line profile was drawn across the wrinkles, and greyscale values (0–255) were extracted for every pixel along the line. Greyscale values were background-corrected and normalized. At least five independent experiments were performed per protein.

Oxidative Cross-Linking of Cysteines in Colony Biofilms.

B. subtilis 48-h grown complex colonies were collected from the agar plate by using a sterile loop and suspended in 250 μL of LB. The biomass was disrupted by passage through a 23 × 1 needle 10 times. The cell suspension obtained was incubated for 15 min at 37 °C with 1.8 mM Cu(II)-(o-phenanthroline)₃ (hereafter CuPhe) or 10 mM DTT, followed by centrifugation and wash of the pellet with 250 μL of PBS. After centrifugation, the pellet was suspended in 250 μL of PBS, followed by a 15 min incubation with 8 mM N-ethylmaleimide/10 mM EDTA to stop the reaction (41, 42). Samples were then centrifuged, and the pellet was washed with 250 μL of PBS. The cell pellet was suspended in 250 µL of BugBuster Master Mix (Novagen), followed by gentle sonication to promote the release of the proteins from the biofilm matrix. The samples were then incubated at room temperature with agitation for 20 min, and the insoluble cell debris was removed by centrifugation at 17,000 × g for 10 min at 4 °C. The proteins samples were then analyzed by Western blot.

Western Blot Analysis.

The 1.5 µg of total protein extract (see above) was separated on a 14% SDS/PAGE before transfer onto PVDF membrane (Millipore) by electroblotting at 25 V for 2 h. The membrane was incubated for 16 h in 3% (wt/vol) powdered milk in TBS [20 mM Tris·HCl (pH 8.0) and 0.15 M NaCl] at 4 °C with shaking. This step was followed by 2-h incubation with purified anti-BslA antibody at a dilution of 1:500 (vol/vol) in TBS in 3% powdered milk wash buffer (TBS + 0.05% Tween 20). The membrane was washed by using wash buffer (TBS + 0.05% Tween 20) and incubated for 45 min with the secondary antibody conjugated to horseradish peroxidize [goat anti-rabbit (Pierce)] at a dilution of 1:5,000. The membrane was washed, developed, and exposed to X-ray film. Western blot analysis for YweA detection was performed as described above with purified anti-YweA antibody at a dilution of 1:10,000 (vol/vol).

Oxygen Profiling of Biofilms.

Overnight cultures of B. subtilis strain 3610 were inoculated from a streaked plate and subsequently grown in LB at 37 °C with shaking at 250 rpm for 12–16 h. Precultures were diluted 10-fold in LB and grown at 37 °C with shaking at 250 rpm until OD_600nm ∼ 1.0. Five microliters of the culture were spotted onto an MSgg agar plate and grown at 30 °C for 2 d before analysis. A 25-µm-tip Clark-type oxygen microsensor (Unisense OX-25) was used to measure the oxygen concentrations. The oxygen microsensor was calibrated according to manufacturer’s instructions, and measurements were taken throughout the depth of the biofilm (step size = 5 µm, measurement period = 3 s, wait time between measurements = 3 s). Four different colonies were probed, and representative data are shown.

Redox Profiling of Biofilms.

Biofilms were grown as described above, and a 25-µm-tip redox microelectrode with an external reference (Unisense RD-25 and REF-RM) was used to measure the extracellular redox potential. After calibrating the redox microelectrode according to the manufacturer’s instructions, redox measurements were taken throughout the depth of the biofilm (step size = 5 µm, measurement period = 3 s, wait time between measurements = 5 s). The redox potential was set to zero at the surface of the colony, and relative values are plotted. Three different colonies were probed, and representative data are shown.

Sporulation Assay.

For heat-resistant spore quantification, colony biofilms were grown for 48 h at 30 °C. Cells were collected in 1 mL of saline solution, disrupted by passage through a 23 × 1 needle 10 times, and subsequently subjected to mild sonication (20% amplitude, 1 s on, 1 s off, for 5 s total) to liberate bacterial cells from the matrix. To kill vegetative cells, the samples were incubated for 20 min at 80 °C. To determine viable cell counts, serial dilutions were plated before and after the 80 °C incubation on LB agar supplemented with 100 μg⋅ml⁻¹ spectinomycin. The percentage of spores was established by colony-forming unit counting, and results are presented as the percentage of colony-forming units obtained after incubation of the samples for 20 min at 80 °C, divided by the number of colony-forming units obtained before the heat inactivation.

Cell Survival upon Chlorhexidine Gluconate Exposure.

To test biofilm resistance to chlorhexidine gluconate, colony biofilms were grown for 48 h at 30 °C. Droplets of 5 µL of 1% (vol/vol) chlorhexidine gluconate were placed on the biofilm surface (near the periphery) for 5 min at room temperature. The solution was removed, and a punch biopsy of 5 mm in diameter was recovered (this area encompassed the entire exposed region). This sample was transferred immediately into 500 µL of saline solution, thus diluting any remaining chlorhexidine gluconate, disrupted by passage through a 23 × 1 needle 10 times and washed once with saline solution. The sample was subsequently subjected to mild sonication (20% amplitude, 1 s on, 1 s off, for 5 s total) to liberate bacterial cells from the matrix. Serial dilutions of the cell suspension were made, and 100 μL was plated onto LB agar supplemented with 100 μg⋅mL⁻¹ spectinomycin; saline solution was used as a control for the process. The percentage survival was evaluated by colony-forming unit counting, and results are presented as the percentage of colony-forming units obtained after exposure to 1% (vol/vol) chlorhexidine gluconate, divided by the number of colony-forming units recovered on the control spots.

Immunofluorescence.

To prepare cross-sections for microscopy, biofilms were grown from strains constitutively expressing GFP for 2 d in standard conditions (Table S1) as previously described (12). A quarter of the colony was excised and placed into optimum cutting temperature compound (Agar Scientific) and frozen in iso-pentene chilled with liquid nitrogen. A Leica CM3050 S cryomicrotome was used to cut cross-sections of this colonies (10–12 μm), which were placed onto SuperFrost Ultra Plus adhesion microscope slides (VWR). The cross-sections were then fixed for 10 min in paraformaldehyde (4% in TBS), washed three times with TBS, and blocked overnight in 2% (wt/vol) fish gelatin (Sigma) prepared in TBS. After washing the cross-sections, anti-BslA antibody diluted 1:200 in AbDil solution (2% BSA and 0.1% azide in TBS) was applied into the cross-sections and incubated for 2.5 h at room temperature. After three washes of 5 min each with TBS, the secondary antibody was applied in a dilution of 1:150 in AbDil and incubated for 90 min at room temperature. The secondary antibody used was DyLight594-conjugated Affinity Pure Donkey Anti-Rabbit IgG (H+L) (Jackson ImmunoResearch). After the incubation, the coverslips were washed 3 times with TBS (5 min each) and mounted in anti-fade containing medium [0.5% p-phenylenediamine (Free base; Sigma), 20 mM Tris (pH 8.8), and 90% glycerol] (43) and sealed with nail varnish. Samples were imaged by using a Zeiss LSM700 confocal scanning laser microscope fitted with 488- and 555-nm lasers and an EC PlanNeofluar 40×/NA 1.30 oil differential interference contrast (DIC) M27 or an alpha Plan-Apochromat 100×/NA 1.49 oil DIC objective. Images were captured and analyzed using Zen2011 software, which was also used to prepare the images presented.