The Bacterial Tyrosine Kinase Activator TkmA Contributes to Biofilm Formation Largely Independently of the Cognate Kinase PtkA in Bacillus subtilis Tantan Gao,a,b,c Jennifer Greenwich,b Yan Li,a Qi Wang,a Yunrong Chaib Department of Plant Pathology, College of Agronomy and Biotechnology, China Agricultural University, Beijing, Chinaa; Department of Biology, Northeastern University, Boston, Massachusetts, USAb; Department of Molecular and Cellular Biology, Harvard University, Cambridge, Massachusetts, USAc

ABSTRACT

In Bacillus subtilis, biosynthesis of exopolysaccharide (EPS), a key biofilm matrix component, is regulated at the posttranslational level by the bacterial tyrosine kinase (BY-kinase) EpsB. EpsB, in turn, relies on the cognate kinase activator EpsA for activation. A concerted role of a second BY-kinase– kinase activator pair, PtkA and TkmA, respectively in biofilm formation was also indicated in previous studies. However, the exact functions of PtkA and TkmA in biofilm formation remain unclear. In this work, we show that the kinase activator TkmA contributes to biofilm formation largely independently of the cognate kinase, PtkA. We further show that the biofilm defect caused by a ⌬tkmA mutation can be rescued by complementation by epsA, suggesting a functional overlap between TkmA and EpsA and providing a possible explanation for the role of TkmA in biofilm formation. We also show that the importance of TkmA in biofilm formation depends largely on medium conditions; the biofilm defect of ⌬tkmA is very severe in the biofilm medium LBGM (lysogenic broth [LB] supplemented with 1% [vol/vol] glycerol and 100 ␮M MnSO4) but marginal in another commonly used biofilm medium, MSgg (5 mM potassium phosphate [pH 7.0], MOPS [100 mM morpholinepropanesulfonic acid] [pH 7.0], 2 mM MgCl2, 700 ␮M CaCl2, 50 ␮M MnCl2, 50 ␮M FeCl3, 1 ␮M ZnCl2, 2 ␮M thiamine, 0.5% glycerol, 0.5% glutamic acid, 50 ␮g/ml tryptophan, 50 ␮g/ml threonine, and 50 ␮g/ml phenylalanine). The molecular basis for the medium dependence is likely due to differential expression of tkmA and epsA in the two different media and complex regulation of these genes by both Spo0A and DegU. Our studies provide genetic evidence for possible cross talk between a BY-kinase activator (TkmA) and a noncognate kinase (EpsB) and an example of how environmental conditions may influence such cross talk in regulating biofilm formation in B. subtilis. IMPORTANCE

In bacteria, biosynthesis of secreted polysaccharides is often regulated by bacterial tyrosine kinases (BY-kinases). BY-kinases, in turn, rely on cognate kinase activators for activation. In this study, we investigated the role of a BY-kinase activator in biofilm formation in Bacillus subtilis. We present evidence that different BY-kinase activators may functionally overlap each other, as well as an example of how activities of the BY-kinase activators may be highly dependent on environmental conditions. Our study broadens the understanding of the complexity of regulation of the BY-kinases/kinase activators and the influence on bacterial cell physiology.

B

iofilms are multicellular communities of bacteria with highly complex structures and distinct morphological features (1–4). One of the most important characteristics of biofilms is the presence of a self-produced extracellular matrix, which allows individual cells within the biofilm to stick to each other (2, 3, 5). In Bacillus subtilis, the biofilm matrix consists of an exopolysaccharide (EPS); the amyloid fiber-like protein TasA; and a small hydrophobin, BslA (6–11). The regulatory circuit that controls the expression of the matrix-encoding genes has been well studied in B. subtilis (5, 12–14). EPS and TasA fibers are produced by the protein products of two matrix operons, epsA-epsO and tapA-sipW-tasA, respectively (8, 15). These two operons are normally repressed by the biofilm repressor SinR and by the transition state regulator AbrB (8, 16–18). Repression by SinR is relieved when, under biofilm induction, a small antirepressor, SinI, is expressed, which antagonizes SinR through direct protein-protein interactions (19–21). The gene sinI is activated by the master regulator Spo0A in response to environmental and cellular signals (22). The gene for the small hydrophobin BslA was shown to be under the control of the response regulator DegU and two tran-

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scription repressors, AbrB and SinR, either directly or indirectly (23). The master regulator Spo0A is positioned at the heart of the genetic network for control of alternative cell fates (motile cells, matrix producers, spore formers, etc.) in B. subtilis (24). Spo0A

Received 2 June 2015 Accepted 12 August 2015 Accepted manuscript posted online 17 August 2015 Citation Gao T, Greenwich J, Li Y, Wang Q, Chai Y. 2015. The bacterial tyrosine kinase activator TkmA contributes to biofilm formation largely independently of the cognate kinase PtkA in Bacillus subtilis. J Bacteriol 197:3421–3432. doi:10.1128/JB.00438-15. Editor: W. W. Metcalf Address correspondence to Qi Wang, [email protected], or Yunrong Chai, [email protected]. T.G. and J.G. contributed equally to this work. Supplemental material for this article may be found at http://dx.doi.org/10.1128 /JB.00438-15. Copyright © 2015, American Society for Microbiology. All Rights Reserved.

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governs endospore formation by regulating hundreds of genes involved in sporulation in B. subtilis (24, 25). Spo0A is also involved in biofilm formation through its control of several regulatory genes, including sinI and abrB (16, 26, 27). The activity of Spo0A is under complex regulation at multiple levels. The Spo0A protein is activated by protein phosphorylation, which depends on the Kin family of sensor histidine kinases (from KinA to KinE) directly, or indirectly via a phosphor relay (mediated by the phosphor transfer proteins Spo0F and Spo0B) (28–30). The histidine kinases sense a variety of environmental or physiological signals and subsequently promote signal transduction, leading to Spo0A activation (13, 31–36). DegU is a global response regulator whose phosphorylation is carried out by its cognate sensor histidine kinase, DegS (37, 38). DegU is involved in motility, chemotaxis, biofilm formation, synthesis of secreted enzymes, and production of ␥-poly-DL-glutamic acid (23, 37, 39, 40). The role of DegU in biofilm formation is quite complex, because DegU controls multiple targets that have been shown to be involved in biofilm formation, both positively and negatively (23, 39, 40). For example, the gene for BslA is under the positive control of DegU (23). In addition to functioning as an activator, high levels of DegU were shown to inhibit the matrix operons, epsA-epsO and tapA-sipW-tasA, although the biological significance of such negative regulation is less well understood (39). Bacterial tyrosine kinases (BY-kinases) belong to a minor family of protein kinases in bacteria (41, 42). They are often associated with the biosynthesis of secreted polysaccharides (43). In Grampositive bacteria, BY-kinases pair with cognate transmembrane kinase activators for activation. The activator protein contains an extracellular domain, which is speculated to sense environmental signals. The activities of BY-kinases also seem to be self-regulated by autophosphorylation at the tyrosine residues. BY-kinases in turn regulate the target proteins by protein phosphorylation. In B. subtilis, EPS biosynthesis is controlled, at the posttranslational level, by the BY-kinase EpsB and its activator, EpsA, in addition to the known regulation of the epsA-epsO operon at the transcriptional level (44, 45). A recent study proposed a feedback mechanism for the regulation of EPS biosynthesis in B. subtilis (46). According to the study, the self-produced exopolysaccharide molecules act as a quorum-sensing-like signal to alter the activity of EpsA, which then interacts with EpsB and stimulates the activity of EpsB (44, 46). EpsB, in turn, regulates the activity of EpsE by protein phosphorylation (46). EpsE is known for its dual function both as a molecular clutch for the control of flagellar motility and as a glycosyltransferase involved in EPS biosynthesis (47, 48). B. subtilis also possesses a second BY-kinase– kinase activator pair, PtkA and TkmA, respectively (45, 49). The genes for the two proteins are located in a presumptive operon with two other genes, ptpZ and ugd (Fig. 1A). The ptpZ gene encodes a phosphatase that counteracts the activity of BY-kinase by dephosphorylating the kinase, as well as the targets of the kinase (45), and ugd encodes a UDP-glucose dehydrogenase (Ugd) (Fig. 1A) (50). The biological functions of TkmA, PtkA, and PtpZ have been investigated in previous studies. It was shown that the protein targets of the PtkA kinase include Ugd and TuaD (involved in teichuronic acid biosynthesis), as well as the single-strand DNA-binding proteins SsbA and SsbB (45, 49, 51). Recent studies also indicated the roles of TkmA and PtkA in biofilm formation. However, the exact

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functions of TkmA and PtkA in biofilm formation remain unclear (44, 52). In this work, we show that TkmA contributes significantly to biofilm formation in B. subtilis, but largely independently of the cognate kinase, PtkA. We provide genetic evidence suggesting that TkmA may functionally overlap EpsA during biofilm formation. Second, we show that the importance of TkmA in biofilm formation is highly medium dependent; the tkmA deletion mutant had a very severe biofilm defect in the biofilm medium LBGM (see below), but the defect was marginal in another commonly used biofilm medium, MSgg. We investigated the molecular basis for the medium dependence effect. MATERIALS AND METHODS Strains, media, and growth conditions. For general purposes, B. subtilis strains PY79 and 3610 and their derivatives were grown at 37°C in lysogenic broth (LB) (10 g tryptone, 5 g yeast extract, and 5 g NaCl per liter broth) or on solid LB medium supplemented with 1.5% agar. For assays of biofilm formation, LBGM or MSgg medium was used. LBGM is composed of LB supplemented with 1% (vol/vol) glycerol and 100 ␮M MnSO4 (36). The recipe for MSgg is as follows: 5 mM potassium phosphate (pH 7.0), 100 mM MOPS (morpholinepropanesulfonic acid) (pH 7.0), 2 mM MgCl2, 700 ␮M CaCl2, 50 ␮M MnCl2, 50 ␮M FeCl3, 1 ␮M ZnCl2, 2 ␮M thiamine, 0.5% glycerol, 0.5% glutamic acid, 50 ␮g/ml tryptophan, 50 ␮g/ml threonine, and 50 ␮g/ml phenylalanine (53). MSgg was supplemented with 1.5% Bacto agar for solid plates. Escherichia coli DH5␣ was used as a host for molecular cloning and was grown at 37°C in LB medium. When required, antibiotics were added at the following concentrations for growth of B. subtilis: 1 ␮g/ml of erythromycin, 5 ␮g/ml of tetracycline, 5 ␮g/ml of chloramphenicol, 100 ␮g/ml of spectinomycin, and 10 ␮g/ml of kanamycin. For growth of E. coli, antibiotics were added at the following concentrations: 100 ␮g/ml of ampicillin, 50 ␮g/ml of kanamycin, and 10 ␮g/ml of tetracycline. The strains and plasmids used in this work are summarized in Tables S1 and S2, respectively, in the supplemental material. DNA manipulation. To extract genomic DNA from overnight cultures of B. subtilis, 1.5 ml cells was harvested in a microcentrifuge tube at 13,000 ⫻ g for 2 min and resuspended in 450 ␮l of distilled water. Fifty microliters of 0.5 M EDTA (pH 8.0) and 60 ␮l freshly prepared lysozyme (20 mg/ml) were added. The mixture was incubated for 30 min at 37°C, followed by addition of 650 ␮l nucleus lysis solution and 250 ␮l protein precipitation solution (Promega). The mixture was rigorously vortexed for 1 min, followed by centrifugation at 15,000 ⫻ g for 5 min; 900 ␮l of the soluble fraction was transferred into a new microcentrifuge tube, and 600 ␮l isopropyl alcohol was added. Centrifugation was repeated at 15,000 ⫻ g for 5 min. The pellet was washed with 70% ethanol and air dried for 30 min before being dissolved in 100 ␮l of nuclease-free water. The general methods for molecular cloning followed published protocols (54). Restriction enzymes (NEB) were used according to the manufacturer’s instructions. Transformations of plasmid DNA into B. subtilis strains were performed as described previously (55). SPP1 phage-mediated transduction was also used to transfer antibiotic-marked DNA fragments among different strains (8, 56). Long-flanking PCR mutagenesis was used to generate insertional deletion mutations (57). The generated PCR constructs were introduced into B. subtilis by genetic transformation and integrated into neutral integration sites (e.g., amyE) in the genome of B. subtilis by double-crossover recombination. The oligonucleotides used in this study are listed in Table S3 in the supplemental material. Strain construction. To construct the insertional deletion mutant of the tkmA-ugd operon (FC72) and the insertional deletion mutants for each of the individual genes in the operon (TG34 [⌬tkmA]), TG35 [⌬ptkA], TG37 [⌬ptpZ], and TG38 [⌬ugd]), long-flanking PCR mutagenesis was applied to generate the mutations. The primers used to generate each of the mutations were as follows: delta-tkmA-P1, delta-tkmA-P2,

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FIG 1 Analyses of the regulatory sequences in the tkmA-ugd operon. (A) Genetic arrangement of the tkmA-ugd operon. Promoters and predicted transcription terminators in the intergenic regions between ywqB and tkmA and between ptkA and ptpZ are indicated by arrows and stem-loops, respectively. A previously characterized small ORF with unknown function (66) in the intergenic region of ywqB and tkmA is highlighted in yellow. The protein products encoded by individual genes in the tkmA-ugd operon are noted below the diagram. BY, bacterial tyrosine. (B) Sequence analysis of the intergenic region between ywqB and tkmA. The ⫺35 and ⫺10 motifs and the transcription start sites (⫹1) of the two ␴A-dependent promoters are annotated, one for the expression of the small ORF highlighted in yellow and the other for the tkmA-ugd operon. The predicted transcription terminator for the small ORF is in purple. Two putative Spo0A⬃P binding sites that perfectly match the consensus sequence (OA box; TGTCGAA) are indicated. The 250-bp DNA sequence used for generating the PtkmA-lacZ reporter fusion is boxed. (C) Sequence analysis of the intergenic region between ptkA and ptpZ. The DNA sequence immediately downstream of ptkA is in purple and folded into a stem-loop structure with the indicated free energy (⌬G). The stop codon of ptkA is in blue, while the start codon of ptpZ is in red.

delta-ugd-P3, and delta-ugd-P4 for ⌬tkmA-ugd; delta-tkmA-P1 to deltatkmA-P4 for ⌬tkmA; delta-ptkA-P1 to delta-ptkA-P4 for ⌬ptkA; deltaptpZ-P1 to delta-ptpZ-P4 for ⌬ptpZ; and delta-ugd-P1 to delta-ugd-P4 for ⌬ugd. The plasmid pDG1515(Tetr) or pAH52(Mlsr) was used as the template during PCR. To construct the reconstituted strains YC997, YC999, and TG43 with a point mutation (G¡A) in the 5= untranslated region of sinR in the native locus, the plasmid pYC192 (20) was used as the template for sinR mutagenesis. pYC192 contains ⬃5 kb of flanking sequence of sinR, spanning from the upstream yqhG, yqhH, and sinI genes to the downstream tasA gene. A kanamycin resistance marker was also inserted into an intergenic region between sinR and tasA. The insertion does not alter the biofilm robustness of the cells (20). Mutagenesis primers PsinR2-F and PsinR2-R were used in the PCR to introduce the point mutation (G¡A) into the 5= untranslated region of sinR in pYC192 (the original sequence was GGAAGGTGATGACATTG; the nucleotide substitution is shown in boldface italics), following application of the site-directed mutagenesis kit (Qiagen). The resulting plasmid, pYC288, was directly introduced into 3610, FC72 (⌬tkmA-ugd), and TG34 (⌬tkmA) by transformation, generating strains YC997, YC999, and TG43, respectively. Successful introduction of the point mutation in the 5= untranslated region of sinR at the native chromosomal locus in the above-mentioned strains was confirmed by PCR amplification of the region and DNA sequencing.

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Construction of the reconstituted strains YC998, YC1200, and TG44 with the point mutation (G¡C) in the 5= untranslated region of sinR in the native locus was similar to that described above, except that the mutagenesis primers PsinR3-F and PsinR3-R were used in the PCR to introduce the point mutation (G¡C) into the 5= untranslated region of sinR in pYC192, following application of the site-directed mutagenesis kit (Qiagen). The resulting plasmid, pYC289, was then directly introduced into 3610, FC72 (⌬tkmA-ugd), and TG34 (⌬tkmA) by transformation, generating strains YC998, YC1200, and TG44. To complement the ⌬tkmA mutation with the wild-type (WT) copy of tkmA, the promoter sequence and the open reading frame (ORF) of tkmA were amplified by PCR, using genomic DNA of B. subtilis 3610 as the template and primers PtkmA-P1 and tkmA-P4. The PCR products were cloned into the integration plasmid pDG1662 (58) by isothermal assembly following a published protocol (59), resulting in the recombinant plasmid pTG05. The recombinant plasmid was then introduced into the B. subtilis laboratory strain PY79 by genetic transformation for the double-crossover recombination of the DNA sequences at the amyE locus. The amyE integration fragment was then introduced into 3610 by SPP1 phage-mediated transduction, generating TG47. To complement the ⌬tkmA mutation with the wild-type copy of epsA, the promoter sequence of tkmA and the coding sequence of epsA were amplified by PCR using the genomic DNA of B. subtilis 3610 as the template and primers PtkmA-P1

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and PtkmA-P2 and primers epsA-P3 and epsA-P4, respectively. The PCR products were cloned into the integration plasmid pDG1662 by isothermal assembly (59), resulting in the recombinant plasmid pTG06. The resulting plasmid was similarly introduced into 3610 as described above, generating TG48. To compare expression of the sinR gene in the wild type and the two reconstituted strains containing either the nucleotide substitution G¡A or G¡C in the untranslated region of sinR, the regulatory sequence and the coding sequence for the first 7 amino acid residues of sinR (followed by a stop codon [see the sequence of the primer PsinR-R5]) were amplified by PCR using primers PsinR-F5 and PsinR-R5, and genomic DNAs of B. subtilis strains 3610, YC997, and YC998, respectively, as the templates. The PCR products were digested with EcoRI and BamHI and cloned into the plasmid pDG268 (60), which carries a chloramphenicol resistance marker and a polylinker upstream of the lacZ gene between two arms of the amyE gene, to make a PsinR-lacZ transcriptional fusion, resulting in the recombinant plasmids pTG01, pTG02, and pTG03, respectively. In this reporter plasmid, translation of lacZ is independent of the translation of the first 7 amino acid residues of sinR. To overexpress epsAB in B. subtilis, the epsAB genes were amplified by PCR using genomic DNA of B. subtilis 3610 as the template and primers epsAB-P1 and epsAB-P2. The PCR product was then cloned into the HindIII and BamHI sites of pDR150, which contains a xylose-inducible xylA promoter (a gift from D. Rudner, Harvard Medical School), generating the recombinant plasmid pTG04. The recombinant plasmid was then introduced into the B. subtilis laboratory strain PY79 by genetic transformation for a double-crossover recombination of the DNA sequences at the amyE locus. The amyE integration fragment was then introduced into strains TG34 and FC72 by SPP1 phage-mediated transduction. To construct the reporter fusion for the polyglutamate biosynthesis gene cluster (ywsCAB-ywtC), the promoter sequence of the operon was amplified by PCR using primers PywsC-F1 and PywsC-R1. The PCR product was digested with HindIII and BamHI and cloned into the vector plasmid pDG268, which was also linearized by HindIII and BamHI. The recombinant plasmid was first introduced into PY79 and then into 3610 in a fashion similar to that described above. Characterization of the suppressor mutation. To characterize the suppressor mutation in the strain YC820, the genomic DNA of YC820 was prepared according to the protocol described above. The genomic DNA was used as the template, and primers PsinR-F1 and sinR-R1 were used to amplify both the regulatory and the coding sequences of the sinR gene. The PCR product was then applied for DNA sequencing (Genewiz). The G-to-A point mutation in the 5= untranslated region of sinR was confirmed by DNA sequencing. Assays of ␤-galactosidase activities. For assays of ␤-galactosidase activities, cells were incubated in MSgg or LBGM at 37°C in a water bath with shaking. One milliliter of culture was collected at each indicated time point after inoculation. The cells were spun down, and the pellets were resuspended in 1 ml Z buffer (40 mM NaH2PO4, 60 mM Na2HPO4, 1 mM MgSO4, 10 mM KCl, and 38 mM ␤-mercaptoethanol) supplemented with 200 ␮l/ml freshly made lysozyme. The resuspensions were incubated at 30°C for 15 min. Reactions were started by adding 200 ␮l of 4 mg/ml ONPG (2-nitrophenyl ␤-D-galactopyranoside) and stopped by adding 500 ␮l of 1 M Na2CO3. Samples were briefly spun down. The optical density at 420 nm (OD420) values of the samples were recorded using a Pharmacia ultraspectrometer 2000. The ␤-galactosidase-specific activity was calculated according to the following formula: (OD420/time ⫻ OD600) ⫻ dilution factor ⫻ 1,000. Assays were conducted at least in duplicate. Assays of pellicle and colony biofilms. B. subtilis cells were grown in LB at 37°C to mid-log phase. For colony formation, 2 ␮l of the cells was spotted onto LBGM medium (or MSgg medium) solidified with 1.5% agar. The plates were incubated at 30°C for 72 h prior to analysis. For pellicle formation, 6 ␮l of the cells was mixed with 6 ml of LBGM broth (or

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FIG 2 Colony and pellicle biofilms formed by mutants with various deletions in the tkmA-ugd operon and complementation strains in LBGM. The strains tested in this assay were the WT (3610); various mutants with either the entire tkmA-ugd operon (⌬tkmA-ugd; FC72) or each of the individual genes in the operon (⌬tkmA/TG34, ⌬ptkA/TG35, ⌬ptpZ/TG37, and ⌬ugd/TG38) deleted; and two complementation strains of ⌬tkmA, one by tkmA (TG47) and the other by epsA (TG48). Cells were inoculated in LBGM in 6-well polyvinyl plates (VWR) or on solidified LBGM agar plates and incubated at 30°C for about 48 h (for pellicles) or 72 h (for colony biofilms). All the scale bars for colony biofilms are approximately 0.3 cm long; all the scale bars for pellicle biofilms are 1 cm long.

MSgg broth) in 6-well plates (VWR). The plates were incubated at 30°C for about 48 h. Images were taken using either a SONY NEX-5 digital camera or a SPOT camera (Diagnostic Instruments).

RESULTS

The deletion mutant for the BY-kinase activator TkmA shows a very severe biofilm defect in LBGM. The exact role of the BYkinase PtkA in biofilm formation in B. subtilis remains elusive for the following reasons. First, the alteration in the biofilm phenotype caused by the ⌬ptkA mutation is somewhat mild, which was shown in previous studies (44, 52) and in this study, as well (Fig. 2; see Fig. S1 in the supplemental material). Second, none of the previously identified protein targets of PtkA seems to play a significant role in biofilm formation, including Ugd (UDP-glucose dehydrogenase), whose gene is located in the same operon with ptkA (Fig. 1A and 2) (52). In most of the previous work aimed at characterizing the functions of TkmA and PtkA in biofilm formation, the commonly used biofilm-inducing medium MSgg was applied in the biofilm assays (53). MSgg is derived from a defined minimal medium originally used in biofilm studies in Pseudomonas aeruginosa with supplementation of 0.5% glycerol and 0.5% glutamic acid (53). We recently formulated a new biofilm-inducing medium, LBGM (LB supplemented with 1% glycerol and 100

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␮M MnSO4), and showed that it promotes strong biofilm formation in B. subtilis (36). We decided to perform biofilm assays of the mutants with either the entire tkmA-ugd operon or each of the individual genes in the operon deleted in this new biofilm medium. Our results show that the mutant with the entire operon deleted (⌬tkmA-ugd) has a very severe defect in the formation of both colony and pellicle biofilms in LBGM (Fig. 2). Interestingly, the deletion mutant for the single tkmA gene (⌬tkmA) also has a severe biofilm defect, nearly identical to that of the operon deletion mutant (Fig. 2). On the other hand, the other three individual deletion mutants (⌬ptkA, ⌬ptpZ, and ⌬ugd) formed biofilms that may be best described as mildly altered in the surface architecture (Fig. 2). We next complemented the ⌬tkmA mutant using the wild-type tkmA gene integrated at the ectopic amyE locus. The complementation strain (⌬tkmA amyE::tkmA) formed robust biofilms that were identical to those of the wild-type cells (Fig. 2). Our results suggest that in the newly formulated biofilm medium LBGM, the kinase activator TkmA plays a major role in biofilm formation and that its role is largely independent of the PtkA kinase or any other protein encoded by genes in the operon. We should point out that in previous studies, it was also shown that even in MSgg, the biofilm phenotype of the ⌬tkmA mutant was somewhat more severe than that of the ⌬ptkA mutant (44, 52). The authors speculated that TkmA might have a PtkA-independent activity in contributing to biofilm formation (44, 52). Our current results not only supported this idea, but suggested further that this PtkA-independent activity of TkmA may be the chief reason for its importance in biofilm formation. In addition, the importance of TkmA in biofilm formation is medium condition dependent, since the biofilm defect caused by the ⌬tkmA mutation was very severe in LBGM (Fig. 2) but marginal in MSgg (see Fig. S1 in the supplemental material). A ⌬tkmA-ugd suppressor mutant forms robust biofilms. Next, we decided to further characterize the putative PtkA-independent activity of TkmA in biofilm formation. We took an unbiased genetic approach by searching for ⌬tkmA-ugd suppressor mutants that are capable of forming robust biofilms again in LBGM. When the ⌬tkmA-ugd mutant was streaked out on an LB agar plate for routine cell proliferation, the colonies demonstrated little surface morphology (see Fig. S2A, top, in the supplemental material). We obtained a spontaneous suppressor mutant that showed quite distinct and robust colony morphology on the same plate (see Fig. S2A in the supplemental material). Upon further characterization, we found that the mutant formed robust colony and pellicle biofilms in LBGM very different from those of the parent strain (Fig. 3B). This indicates that the putative suppressor mutation fully rescued the biofilm defect caused by the ⌬tkmAugd mutation. The suppressor mutant also showed extensive cell chains in shaking culture, again, very different from the parent strain (see Fig. S2B in the supplemental material). Based on observations in our previous studies (61, 62), in which spontaneous mutants bearing point mutations in either the sinR or abrB gene were shown to form both robust biofilms and extensive cell chains, we decided to map the suppressor mutation by trying the targeted approach first and sequencing those biofilm-related regulatory genes in the suppressor mutant. Interestingly, our sequencing results indeed revealed a single nucleotide substitution in a region immediately upstream of sinR. The mutation (G¡A) (Fig. 3A) seems to be located in the 5= untranslated

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FIG 3 Characterization and reconstitution of the suppressor mutation. (A) The suppressor mutation in YC820 was characterized as a single nucleotide substitution (from G to A) in the 5= untranslated region of sinR. The ⫺35 and ⫺10 motifs and the transcription start site (⫹1) of the promoter of sinR and the single nucleotide change (from G to A) are indicated. The Shine-Dalgarno sequence is in blue, and the start codon (TTG) of sinR is in red. (B) Colony and pellicle biofilm phenotypes in LBGM of the wild type (3610), the tkmA-ugd deletion mutant (FC72), the suppressor mutant of ⌬tkmA-ugd (YC820), and two reconstituted strains that contain the single nucleotide substitution G to A (YC997) or G to C (YC998) in the regulatory region of sinR. All the bars for colony biofilms are approximately 0.3 cm long; all the bars for pellicle biofilms are 1 cm long. (C) Assays for ␤-galactosidase activity were performed to compare the activities of the strains harboring the transcriptional fusion of either the wild-type sinR gene (TG40) (diamonds) or the sinR alleles with either a G-to-A (TG41; squares) or a G-to-C (TG42; triangles) substitution in the regulatory region of sinR to the lacZ gene. The reporter fusion was integrated at the chromosomal amyE locus of a ⌬epsH mutant (RL4548). (D) Assays of ␤-galactosidase activity were performed to compare the activities of the strains that harbor the reporter fusion for the epsA-epsO operon (PepsA-lacZ) integrated at the amyE locus and contain either the wild-type sinR gene at the native locus (YC952; diamonds) or the sinR mutant alleles with either a G-to-A (YC1201; squares) or a G-to-C (YC1202; triangles) change in the 5= untranslated region of sinR. The results are representative of three independent assays.

region of sinR between the transcription start site (⫹1, indicated by the bent arrow) and the translation start site (TTG), next to the Shine-Dalgarno-like sequence (5=-GGAAGG-3=), for ribosome binding. To confirm that the single nucleotide change in the 5= untranslated region of sinR is solely responsible for the suppressor phenotype, we reconstituted the point mutation on the chromosome of the ⌬tkmA-ugd mutant. To do so, we introduced two different single nucleotide changes (G to A, as in the original suppressor mutant, as well as G to C) into the 5= untranslated region in the native sinR locus, following a previously published protocol (62) (see Materials and Methods). We then tested the biofilm pheno-

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types of the two reconstituted strains. Both strains formed robust colony and pellicle biofilms in LBGM identical to that of the original suppressor mutant (Fig. 3B). This result confirms that the single nucleotide change in the 5= untranslated region of sinR is solely responsible for the suppression of the biofilm defect caused by the ⌬tkmA-ugd mutation. The reconstituted mutations also completely rescued the biofilm defect by the single ⌬tkmA deletion mutation (data not shown). The suppressor mutation lowers sinR expression. Since the suppressor mutation is located in the 5= untranslated region of sinR, we suspected that the mutation might alter sinR expression at the posttranscriptional level by affecting either the mRNA stability or translation initiation of sinR. To test the first possibility, we created a transcriptional reporter by fusing the regulatory region and the coding sequence for the first 7 amino acids of sinR (followed by addition of a stop codon) to the lacZ gene. Since this fusion contains a separate ribosome binding site and the translation start site for the lacZ gene, it is used to report the mRNA abundance of the lacZ transcript starting from the untranslated region of sinR. In parallel, we also introduced the same point mutations (either G to A or G to C) into the 5= untranslated region of sinR in the reporter fusion (see Materials and Methods). B. subtilis 3610 derivatives bearing either the wild-type reporter or the reporter fusion with the point mutation were assayed for ␤-galactosidase activities. The results (Fig. 3C) showed that the two reporter strains harboring the point mutation (TG41 and TG42 [squares and triangles, respectively]) had lower activities than the wild-type reporter strain (TG40 [diamonds]), an indication that the suppressor mutation negatively impacted sinR expression, possibly by affecting the mRNA stability of sinR. In theory, it is also possible that the mutation impacts both mRNA stability and translation initiation of sinR (e.g., by also influencing ribosome binding). We chose not to test the latter possibility, since the putative change in mRNA stability is sufficient to explain what we have observed. How the point mutations in the 5= untranslated region of sinR may affect sinR mRNA stability is still unclear to us. A previous study showed that the mRNA stability of sinR was regulated by specific RNA nucleases, whose activities significantly affect biofilm formation in B. subtilis (63). Thus, we speculated that sinR could be one of the prime targets for mRNA stability control in B. subtilis. epsA complementation rescued the biofilm defect caused by ⌬tkmA. Our results suggest that lowered expression of sinR may be the cause of the suppressor phenotype. This may in turn be due to overexpression of some of the SinR-repressed genes, since SinR acts as a biofilm master repressor. The SinR regulon has been well characterized, including the tapA-sipW-tasA operon for production and deployment of TasA protein fibers, the epsA-epsO operon for biosynthesis of EPS, and the lutABC operon for lactate utilization (8, 15, 64). We decided to focus on the epsA-epsO operon, partly because epsAB, the first two genes in the operon, are homologous to tkmA-ptkA. To test whether epsA-epsO is overexpressed in the suppressor mutant, we introduced a transcriptional reporter for the epsA-epsO operon (PepsA-lacZ) into the strains with either the wild-type sinR or the sinR mutant allele at the native locus that we constructed previously, resulting in strains YC952 (WT), YC1201 (G¡A), and YC1202 (G¡C), respectively. We compared the ␤-galactosidase activities of the above-mentioned three strains. To avoid cell clumping during shaking growth due to hyperactivity of SinR in some of the strains (8), an ⌬epsH mutation was introduced into the reporter strains. Our results showed

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FIG 4 Overexpression of epsAB rescues the biofilm defect caused by ⌬tkmA. The epsAB genes were placed under the control of the xylose-inducible promoter (PxylA-epsAB), and the fusion was introduced into the ⌬tkmA-ugd mutant and the ⌬tkmA mutant, resulting in TG45 and TG48, respectively. In the presence of xylose (0.1% [vol/vol]), both strains were able to form robust colony (A) and pellicle (B) biofilms in LBGM comparable to that of the wildtype cells, whereas in the absence of xylose, no strong biofilm was observed in the engineered strains. All the bars for colony biofilms are approximately 0.3 cm long; all the bars for pellicle biofilms are 1 cm long. OE, overexpression.

that the two strains with the point mutation in the 5= untranslated region of sinR (YC1201 and YC1202 [Fig. 3D, squares and triangles]) had much higher activities than the wild-type strain (YC952 [diamonds]). The reason why a mild decrease in the expression of sinR may lead to such a drastic increase in the expression of epsAepsO is due to hypersensitivity of SinR-mediated repression, a unique feature of SinR activity characterized in our previous studies (20, 62). We speculated that overexpression of epsA-epsO (or, more specifically, epsAB) in the suppressor mutant may compensate for the loss of tkmA. To test our hypothesis, we created an inducible epsAB construct by fusing epsAB to a xylose-inducible promoter (PxlyA-epsAB) (see Materials and Methods) and integrated the fusion into the chromosomal amyE locus in the ⌬tkmA-ugd and ⌬tkmA mutants. We then tested biofilm formation by those engineered strains in LBGM in the absence or presence of xylose. Interestingly, in the presence of xylose, the biofilm defects in both the ⌬tkmA-ugd and the ⌬tkmA mutants were largely rescued (Fig. 4), whereas in the absence of xylose, no suppression of the biofilm defect was seen. We conclude that loss of tkmA (or the entire tkmA-ugd operon) can be fully compensated for by overexpression of epsAB. To further narrow down the role of compensation to either epsA or epsB, we repeated the complementation experiment for the ⌬tkmA mutant. This time, we placed the epsA gene under the control of the tkmA promoter (PtkmA-epsA) and integrated the engineered construct into the amyE locus of the ⌬tkmA mutant. We similarly tested biofilm formation by the strain in LBGM. As

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shown in Fig. 2, this complementation strain formed fairly robust colony and pellicle biofilms, indicating that epsA itself (at a comparable expression level) is able to compensate for the loss of tkmA. It also implies that the PtkA-independent activity of TkmA in biofilm formation may have to do with the ability of TkmA to complement EpsA activities. If so, it is conceivable that TkmA may cross talk with EpsB, the cognate BY-kinase of EpsA, to regulate EPS biosynthesis. Expression of tkmA is higher in LBGM than in MSgg, while the opposite is true for epsA. Based on our genetic evidence, we propose that the PtkA-independent activity of TkmA when contributing to biofilm formation is likely due to its ability to complement EpsA activities, but it is still unclear to us why the importance of tkmA in biofilm formation differs significantly in the two different biofilm media (LBGM versus MSgg). One possible scenario is that the tkmA gene is differentially expressed in the two different media, being higher in LBGM and lower in MSgg. Higher expression of tkmA presumably leads to proportionally more significant contributions of TkmA to EPS biosynthesis and biofilm formation in LBGM than in MSgg. To test our hypothesis, we constructed a transcriptional reporter fusion for tkmA by amplifying and fusing the promoter sequence of tkmA (from about 250 bp upstream to the start of tkmA [Fig. 1]) to lacZ. We then integrated this reporter (PtkmA-lacZ) into the chromosomal amyE locus of B. subtilis 3610 and conducted a bioassay comparing the expression levels of tkmA in LBGM and MSgg by using the engineered B. subtilis reporter strain (TG49). Our results show that the expression of tkmA increased over time in both media (Fig. 5A). Interestingly, the increase was clearly greater in LBGM (Fig. 5A, squares) than in MSgg (Fig. 5A, diamonds). This result indicates that tkmA is more highly expressed in LBGM than in MSgg. If TkmA were to compete with EpsA for cross talking with EpsB, then the expression of epsA in LBGM and MSgg might also matter. We applied a similar approach to test whether there is differential expression of epsA in the two different media. The B. subtilis strain bearing the PepsA-lacZ transcriptional fusion (YC110 [65]) was cultured in LBGM and MSgg and assayed for ␤-galactosidase activities. Interestingly, and opposite to what was seen for the PtkmA-lacZ reporter, cells harboring the PepsA-lacZ reporter showed much higher activities in MSgg (Fig. 5B, diamonds) than in LBGM (Fig. 5B, squares). Taken together, we conclude that tkmA is expressed at a higher level in LBGM than in MSgg, whereas for epsA, it is expressed at a lower level in LBGM than in MSgg. An inference from this is that in LBGM, TkmA may compete more effectively with EpsA for interaction with EpsB and thereby contribute more to the regulation of EPS biosynthesis and biofilm formation than in MSgg. If this is true, it may explain why the importance of TkmA in biofilm formation differs significantly between LBGM and MSgg. tkmA and epsA are regulated by Spo0A and DegU in a complex fashion. To further understand what caused the differential expression of tkmA and epsA in LBGM and MSgg, we decided to revisit the transcriptional regulation of tkmA and epsA in the two different biofilm media. Transcriptional regulation of the tkmAugd and the epsA-epsO operons has been investigated in previous studies (25, 40, 52). The results from microarray analysis suggest that the tkmA-ugd operon belongs to the Spo0A regulon (25). It was also shown that phosphorylated Spo0A (Spo0A⬃P) bound to the regulatory sequence of the tkmA-ugd operon in gel mobility shift assays, indicating direct regulation of the operon by Spo0A

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FIG 5 Genetic evidence for differential regulation of tkmA and epsA in LBGM

and MSgg. Shown are assays of the ␤-galactosidase activities of the wild-type strain harboring one of the following fusions: PtkmA-lacZ (TG49) (A), PepsAlacZ (YC110) (B), PywsC-lacZ (YC1275) (C), PflaB-lacZ (YC605) (D), PabrB-lacZ (FC287) (E), or PsinI-lacZ (YC127) (F). The cells were grown in either LBGM (squares) or MSgg (diamonds) in shaking culture. Cell samples were periodically collected and assayed for ␤-galactosidase activity. The error bars represent standard deviations from triplicate assays.

(25). However, that result may need further examination, as we elaborate below. Our analysis of the regulatory region of tkmA (the intergenic sequences between tkmA and ywqB) revealed two putative Spo0A⬃P binding sites that perfectly match the consensus sequence of the so-called OA box (TGTCGAA) (Fig. 1B). The first binding site is located upstream of a ␴A-dependent promoter that drives expression of a putative small open reading frame with unknown function (highlighted in yellow in Fig. 1A and B) (66). The second putative binding site of Spo0A⬃P is located upstream of a ␴A-dependent promoter that possibly drives the transcription of the tkmA-ugd operon (Fig. 1A and B). To test whether Spo0A activates transcription of the tkmA-ugd operon from the second putative OA box, we introduced the previously constructed PtkmAlacZ reporter fusion (note that this reporter fusion contains only the intergenic region between tkmA and the small ORF) into the wild-type strain and the spo0A mutant and compared the ␤-galactosidase activities of the two strains. As shown in Fig. 6A, in the spo0A mutant, the PtkmA-lacZ fusion was expressed at a much lower level, confirming that Spo0A positively regulates the tkmAugd operon, likely by binding to the second OA box. DegU was also shown to positively regulate the tkmA-ugd operon in previous

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FIG 6 Genetic regulation of the tkmA-ugd operon. (A to C) Assays of the ␤-galactosidase activities of the wild-type strain, the spo0A mutant, and the degSU mutant that contain a transcriptional lacZ reporter fusion with the regulatory region of either tkmA (PtkmA-lacZ) (A), ptpZ (PptpZ-lacZ) (B), or epsA (PepsA-lacZ) (C). All the fusions were integrated at the chromosomal amyE loci of the cells. The cells were grown in LBGM with shaking to an OD600 of 1.0 before being harvested for the assays. The error bars represent standard deviations from triplicate assays. (D) Schematic summary of regulation of the tkmA-ugd and epsA-epsO operons by Spo0A and DegU. Spo0A positively regulates both the tkmA-ugd and the epsA-epsO operons, whereas DegU has opposite regulation of the two operons, acting positively on tkmA-ugd but negatively on epsAepsO. DegU also negatively regulates the expression of ptpZ (and possibly ugd) from a putative internal promoter just upstream of ptpZ by an unknown mechanism. ⫹, positive regulation; ⫺, negative regulation.

studies (40, 52). We also confirmed that the activity of the same PtkmA-lacZ reporter fusion decreased to very low levels in the degSU mutant compared to that in the wild-type cells (Fig. 6A). Thus, the tkmA gene is under strong positive regulation by both DegU and Spo0A. Further sequence analysis of the operon revealed that between the ptkA and ptpZ genes there lies a 52-bp intergenic region (Fig. 1A and C). A stem-loop-like structure can be predicted based on DNA sequences in this intergenic region, which could function as a putative transcriptional attenuator (Fig. 1C). If this is true, we wondered if there might be a second promoter in the intergenic region driving the expression of just the downstream ptpZ and ugd genes. To test this, we constructed a transcriptional reporter by fusing the 52-bp intergenic region with lacZ, creating PptpZ-lacZ, and introduced this reporter into the wild-type strain and the ⌬spo0A and ⌬degSU mutants for integration at the amyE locus. Wild-type cells harboring the reporter were assayed for ␤-galactosidase activities and showed very little activity in shaking culture, indicating that the the putative internal promoter (if it exists) is not being actively transcribed under normal conditions (Fig. 6B). The same was seen in the spo0A mutant. Surprisingly, the reporter showed drastically increased activities when assayed in the degSU mutant (Fig. 6B), suggesting that DegU strongly represses the phosphatase gene (ptpZ) from the putative internal promoter while simultaneously positively regulating the genes for

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the kinase and the kinase activator (ptkA and tkmA) from the upstream promoter. So, why do cells have such complex regulation of this operon by DegU? We presumed that in the DegU⬃PHIGH cells, not only is the kinase PtkA highly expressed but also, simultaneously, the cells ensure that the production of the counteracting phosphatase, PtpZ, is low due to repression by DegU⬃P, as well as possible transcriptional attenuation immediately after ptkA. Together, these would allow strong activation of the kinase by autophosphorylation due to limited counteracting phosphatase activities, as well as strong activation of the target proteins. For the epsA-epsO operon, previous studies showed that it is regulated by both Spo0A and DegU (8, 39). Spo0A positively and indirectly regulates the operon through its regulation of the sinI and abrB genes (17, 18, 22). A recent study showed that DegU negatively regulates the epsA-epsO operon (39). This was somewhat unexpected, given that DegU is a positive regulator of biofilm formation in B. subtilis. By application of the PepsA-lacZ reporter, we confirmed the positive regulation of epsA-epsO by Spo0A and strong negative regulation of the operon by DegU (Fig. 6C) (39). In summary, the epsA-epsO and the tkmA-ugd operons are under complex regulation by both Spo0A and DegU. We provide a simple scheme to summarize the concerted regulation of the two operons (Fig. 6D). We predict that rising Spo0A activities will increase the production of both TkmA and EpsA. In contrast, rising DegU activities may oppositely affect the levels of the two activator proteins, strongly increasing TkmA expression while holding back the production of EpsA. If so, it is conceivable that environmental conditions favoring high DegU activities will render the role of TkmA more important in biofilm formation. Varied activities of Spo0A and DegU in LBGM and MSgg are responsible for the differential expression of tkmA and epsA. Spo0A and DegU are two well-studied examples of regulatory proteins whose activities are highly dependent on environmental conditions, and both of them regulate the epsA-epsO and the tkmAugd operons. Thus, it is possible that differential expression of tkmA and epsA is a consequence of varied activities of Spo0A and DegU under the two different medium conditions. Here, we focus on the possible role of DegU in the differential expression of epsA and tkmA, whereas the corresponding role of Spo0A is discussed below. We hypothesized that the activities of DegU may be higher in LBGM than in MSgg, which in turn causes higher expression of tkmA but lower expression of epsA in LBGM than in MSgg (Fig. 5A and B). To test our hypothesis, we constructed two transcriptional reporter fusions, PywsC-lacZ and PflaB-lacZ. The first reporter, PywsC-lacZ, allowed us to measure the expression of the ywsCABywtC operon, whose gene products are involved in production of ␥-poly-DL-glutamic acid (67). The PflaB-lacZ transcriptional fusion reports the activity of the fla-che operon, whose gene products are involved in motility and chemotaxis (40). Both operons are known to be positively regulated by DegU (40, 68). The ␤-galactosidase activities of the cells harboring the reporters grown in LBGM and MSgg were assayed. Our results showed that the expression of both reporters was higher in LBGM (Fig. 5C and D, squares) than in MSgg (diamonds), indicating that the activity of DegU varies in the two different media and is likely higher in LBGM than in MSgg. As a reminder, higher DegU activities may oppositely affect the levels of the two activator proteins, strongly

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increasing TkmA expression while repressing the production of EpsA. For the role of Spo0A in the differential expression of tkmA and epsA in the two different media, we similarly hypothesized that Spo0A activities might be higher in LBGM than in MSgg. We took a similar genetic approach to compare the activities of Spo0A in LBGM and MSgg by applying a transcriptional reporter, PabrBlacZ. The abrB gene is known to be directly and negatively regulated by Spo0A. Thus, expression of abrB inversely correlates with Spo0A activities (27). Cells bearing the PabrB-lacZ reporter were grown in LBGM and MSgg and assayed for ␤-galactosidase activity. Our results show that the reporter strain had consistently lower activity when grown in LBGM (Fig. 5E, squares) than in MSgg (diamonds), indicating that Spo0A activities also differ in LBGM and MSgg and were higher in LBGM than in MSgg. Similar to those of DegU, higher Spo0A activities likely also contribute to higher tkmA expression in LBGM than in MSgg. Since Spo0A is also a positive regulator of the epsA-epsO operon (Fig. 6C), one may intuitively predict that higher activities of Spo0A will lead to higher expression of epsA-epsO in LBGM than in MSgg. Spo0A indirectly regulates epsA-epsO, in part through its regulation of sinI (20). SinI is a small antirepressor that antagonizes the master biofilm repressor SinR, leading to derepression of SinR-controlled genes (8, 19). Surprisingly, when we compared the ␤-galactosidase activities from the cells harboring the PsinIlacZ reporter in LBGM and MSgg, the activities of the reporter strain were mildly lower in LBGM (Fig. 5F, squares) than in MSgg (diamonds,). This indicates that, for sinI (and likely SinR-repressed epsA-epsO), higher activities of Spo0A alone do not result in higher expression of those genes in LBGM than in MSgg. An explanation for the above-mentioned result may lie in the regulatory region of sinI, which contains a unique arrangement of both the activator and multiple operator sequences of Spo0A (20). Our previous study showed that optimal expression of sinI depends on intermediate levels of Spo0A, which preferentially binds to the high-affinity activator site but not the low-affinity operator sites. Higher levels of Spo0A instead curtail sinI activation by binding to both the activator and operator sites (20). In toto, we propose that differential expression of tkmA and epsA in the two different biofilm media may be the reason why the importance of TkmA in biofilm formation is highly medium dependent, and this is contributed by both Spo0A and DegU, both of which showed higher activities in LBGM than in MSgg. DISCUSSION

BY-kinases play important roles in the biosynthesis of secreted polysaccharides in bacteria. In B. subtilis, the BY-kinase and the kinase activators, EpsB and EpsA, are involved in the regulation of EPS biosynthesis and biofilm formation (44, 46, 49). Previous studies also suggested that the second BY-kinase PtkA, augments the role of EpsB in biofilm formation (44, 52). However, the exact functions of PtkA and TkmA in biofilm formation remain unclear. In this study, thanks to the application of the new biofilminducing medium LBGM, we show that the mutant for the kinase activator, TkmA, has a severe biofilm defect in LBGM and that the importance of TkmA in biofilm formation is largely independent of the cognate BY-kinase PtkA. Based on our genetic evidence, we postulated that the role of TkmA in biofilm formation might have more to do with shared regulation with EpsA of EpsB. Interestingly, while we were working on this project, a recent study inves-

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tigating cross-regulation among different types of kinases in B. subtilis provided molecular evidence to support our hypothesis (69). Using yeast and bacterial two-hybrid assays, strong interactions between TkmA and EpsB and between TkmA and PtkA but not between EpsA and PtkA were observed in that study (69). In terms of the biological significance of such interactions (cross talk between the noncognate TkmA and EpsB), one simple projection is that EPS biosynthesis and biofilm formation may be regulated at the posttranslational level in response to a broader range of environmental stimuli. In addition, recent studies also showed broad cross-phosphorylation among BY-kinases and other types of kinases (e.g., Ser/Thr kinases), as well as overlapping activities among different pairs of BY-kinases and the kinase activators. All these studies imply that the complex regulation and biological significance of the BY-kinases may have been underestimated (69). Here, we presented a schematic model to summarize our hypothesis (see Fig. S3 in the supplemental material). In that model, EpsA is indicated as sensing self-produced exopolysaccharide molecules in a quorum-sensing-like mechanism (46) while TkmA is speculated to sense unknown environmental signals. It will be interesting to know what signals TkmA senses and how these signals may be physiologically relevant to biofilm formation in future studies. Another interesting observation in this study is that the importance of TkmA in biofilm formation is highly medium dependent. Our evidence suggests that the medium dependence effect may be due to differential expression of tkmA and epsA in LBGM and MSgg. We further propose that differential expression of tkmA and epsA in the two different media is a consequence of complex regulation of the genes by Spo0A and DegU, both of which showed higher activities in LBGM than in MSgg. A model to explain the molecular basis for the medium condition dependence of the importance of TkmA in biofilm formation is shown in Fig. 7. Our study illustrated an interesting example of how medium (or environmental) conditions can significantly alter the activity of the BY-kinase activators in the regulation of the bacterial multicellular development process. What exactly differs between the two different biofilm media that subsequently results in varied activities of both Spo0A and DegU is not known. It is also somewhat difficult to directly compare the two media (one [LBGM] is a rich medium without defined chemical ingredients, and the other [MSgg] is a defined minimal medium). However, it is worth noting that manganese was shown to be critical for Spo0A activities (70). Therefore, larger amounts of manganese in LBGM (100 ␮M) than in MSgg (50 ␮M) may be partially responsible for the difference in Spo0A activities in the two different media. In fact, when manganese was provided at 50 ␮M, the biofilm-inducing activity of the modified LBGM was notably weaker (data not shown). Lastly, we cannot rule out the possibility that LBGM contains an unknown signal at an abundant level for optimal activation of TkmA whereas the putative signal is largely absent or much less abundant in MSgg. The idea of an unknown signal for activation of TkmA is based on the observation that the kinase activator contains an extracellular domain for potential signal sensing, and the presence of such a putative signal is thought to be important for the activity of the kinase activator in interacting with and activating the kinase. Finally, our study also indicates that DegU is not simply a positive regulator of the tkmA-ugd operon; rather, DegU acts as a switch for the kinase/phosphatase (PtkA/PtpZ) activities. We

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FIG 7 Proposed model for the molecular basis of the medium dependence effect in biofilm formation. Based on genetic evidence presented in this study, two different scenarios corresponding to the two different medium conditions (LBGM [A] and MSgg [B]) are shown. For Spo0A, we propose that the levels of Spo0A are higher in LBGM (⫹⫹⫹⫹) than in MSgg (⫹⫹). Consequently, Spo0A directly activates the tkmA-ugd operon at higher levels in LBGM (⫹⫹⫹⫹) than in MSgg (⫹⫹). In contrast, higher levels of Spo0A do not necessarily lead to higher expression of sinI due to the presence of both activator and operator sites of Spo0A in the regulatory region of sinI (20). Thus, Spo0A alone is not expected to cause differential expression of epsA-epsO in LBGM and MSgg. For DegU, we also predict higher activities in LBGM (⫹⫹⫹⫹) than in MSgg (⫹⫹), which lead to higher expression of tkmA-ugd and lower expression of epsA-epsO due to strong repression of epsA-epsO by DegU in LBGM. This model predicts that in LBGM, the TkmA/EspA ratio is much higher than that in MSgg. Therefore, TkmA makes significantly more contributions to the regulation of EPS biosynthesis and biofilm formation by cross-talking with EpsB.

showed that, in parallel with the positive regulation of ptkA from the upstream promoter, DegU also seems to negatively regulate the expression of ptpZ and ugd from a putative internal promoter located between the ptkA and ptpZ genes (Fig. 1A and C). In addition, a putative transcriptional attenuator immediately downstream of ptkA may further reduce the expression of the phosphatase gene (Fig. 1C). Therefore, when DegU activities are high, DegU preferentially promotes expression of ptkA for the kinase activity while simultaneously repressing ptpZ for the phosphatase activity. When DegU activities are low, it derepresses expression of the phosphatase gene from the putative internal promoter. We hope to show that such an intergenic region between the gene for the kinase and the gene for the phosphatase and the transcription attenuator-like stem-loop structure within the intergenic region seem to be fairly conserved in homologous gene clusters in other Bacillus species (see Fig. S4 in the supplemental material). ACKNOWLEDGMENTS We thank Alex Elsholz and Sarah Walker (Harvard University) for advice and helpful discussions during this work. We are also grateful to Rich Losick for his support and mentorship of T. Gao. T. Gao was supported by a fellowship from the China Scholarship Council (201206350144) to study abroad at Harvard University and Northeastern University. Funding for this work was provided by a startup grant from Northeastern University to Y. Chai. It was also partly supported by grants from the National Science Foundation Council (China) (no. 31171893 and 31101480).

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REFERENCES 1. Kolter R, Greenberg EP. 2006. Microbial sciences: the superficial life of microbes. Nature 441:300 –302. http://dx.doi.org/10.1038/441300a. 2. Lemon KP, Earl AM, Vlamakis HC, Aguilar C, Kolter R. 2008. Biofilm development with an emphasis on Bacillus subtilis. Curr Top Microbiol Immunol 322:1–16. 3. O’Toole G, Kaplan HB. 2000. Biofilm formation as microbial development. Annu Rev Microbiol 54:49 – 80. http://dx.doi.org/10.1146/annurev .micro.54.1.49. 4. Stoodley P, Sauer K, Davies DG, Costerton JW. 2002. Biofilms as complex differentiated communites. Annu Rev Microbiol 56:187–209. http://dx.doi.org/10.1146/annurev.micro.56.012302.160705. 5. Vlamakis H, Chai Y, Beauregard P, Losick R, Kolter R. 2013. Sticking together: building a biofilm the Bacillus subtilis way. Nat Rev Microbiol 11:157–168. http://dx.doi.org/10.1038/nrmicro2960. 6. Kobayashi K, Iwano M. 2012. BslA(YuaB) forms a hydrophobic layer on the surface of Bacillus subtilis biofilms. Mol Microbiol 85:51– 66. http://dx .doi.org/10.1111/j.1365-2958.2012.08094.x. 7. Hobley L, Ostrowski A, Rao FV, Bromley KM, Porter M, Prescott AR, MacPhee CE, van Aalten DMF, Stanley-Wall NR. 2013. BslA is a selfassembling bacterial hydrophobin that coats the Bacillus subtilis biofilm. Proc Natl Acad Sci U S A 110:13600 –13605. http://dx.doi.org/10.1073 /pnas.1306390110. 8. Kearns DB, Chu F, Branda SS, Kolter R, Losick R. 2005. A master regulator for biofilm formation by Bacillus subtilis. Mol Microbiol 55: 739 –749. 9. Branda SS, Chu F, Kearns DB, Losick R, Kolter R. 2006. A major protein component of the Bacillus subtilis biofilm matrix. Mol Microbiol 59: 1229 –1238. http://dx.doi.org/10.1111/j.1365-2958.2005.05020.x. 10. Romero D, Aguilar C, Losick R, Kolter R. 2010. Amyloid fibers provide structural integrity to Bacillus subtilis biofilms. Proc Natl Acad Sci U S A 107:2230 –2234. http://dx.doi.org/10.1073/pnas.0910560107.

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