Bacillus subtilis biofilm induction by plant polysaccharides

Significance

The plant growth-promoting bacterium Bacillus subtilis is frequently found associated with plant roots where it protects plants from infection. Here, we demonstrate that B. subtilis root attachment depends on production of an extracellular matrix that holds the cells together in multicellular communities termed biofilms. We found that plant polysaccharides (major components of the plant’s cell wall) act as an environmental cue that triggers biofilm formation by the bacterium. Furthermore, these plant polysaccharides can serve as a carbon source used to produce the extracellular matrix. This work sheds light on how plants stimulate their colonization by this plant growth-promoting rhizobacterium.

Abstract

Bacillus subtilis is a plant-beneficial Gram-positive bacterium widely used as a biofertilizer. However, relatively little is known regarding the molecular processes underlying this bacterium's ability to colonize roots. In contrast, much is known about how this bacterium forms matrix-enclosed multicellular communities (biofilms) in vitro. Here, we show that, when B. subtilis colonizes Arabidopsis thaliana roots it forms biofilms that depend on the same matrix genes required in vitro. B. subtilis biofilm formation was triggered by certain plant polysaccharides. These polysaccharides served as a signal for biofilm formation transduced via the kinases controlling the phosphorylation state of the master regulator Spo0A. In addition, plant polysaccharides are used as a source of sugars for the synthesis of the matrix exopolysaccharide. The bacterium's response to plant polysaccharides was observed across several different strains of the species, some of which are known to have beneficial effects on plants. These observations provide evidence that biofilm genes are crucial for Arabidopsis root colonization by B. subtilis and provide insights into how matrix synthesis may be triggered by this plant.

The rhizosphere, the portion of soil surrounding plant roots, was described over 100 y ago as “a nesting site for a rich and dynamic community of microorganisms” (1). Some of these microorganisms are plant pathogens, whereas others live with plants as mutualists (reviewed in refs. 2–5). Bacteria that thrive in the rhizosphere, colonize the roots, and promote plant growth are known as plant growth-promoting rhizobacteria (6). One such plant growth-promoting rhizobacterium is the Gram-positive bacterium Bacillus subtilis. Widely known as a soil-dwelling bacterium, B. subtilis is also found in association with roots of many different plants (7–9). Because it exhibits many beneficial activities for the plant, B. subtilis is widely used as a biofertilizer (10). In artificial medium, B. subtilis promotes plant growth via the secretion of cytokinin hormones and volatiles that modify plant hormone homeostasis (11, 12). The same volatiles also help the plant cope with salt stress by modulating expression of the high affinity ion transporter HKT1 (13). In addition, B. subtilis can directly prevent infection of the plant by bacterial pathogens by releasing the AiiA enzyme. AiiA is a lactonase that functions by inactivating acyl-homoserine-lactone molecules that regulate the expression of virulence genes by many plant pathogens (14). B. subtilis also secretes surfactin that acts as an antimicrobial against pathogens such as Pseudomonas syringae (15). Finally, B. subtilis can indirectly protect plants by inducing systemic resistance, which affords the plant protection against a broad range of pathogens (16–19).

Plant growth-promoting bacteria are thought to form biofilms on roots (20). Biofilms are assemblages of cells embedded in a matrix composed of exopolysaccharides (EPSs), proteins, and sometimes DNA (reviewed in refs. 21 and 22). Matrix production results in the formation of complex architectures typical of biofilms (23, 24). In B. subtilis, the extracellular matrix is composed of two major components, an EPS and the protein TasA, which polymerizes into amyloid-like fibers (25, 26). Expression of the epsA-O operon, encoding the enzymes involved in EPS production, and tasA is indirectly under the control of the master transcriptional regulator Spo0A (Fig. S1). Spo0A activity depends on its phosphorylation state (27). Spo0A phosphorylation is controlled by five histidine kinases (KinA–KinE), which respond to different environmental cues (28). Phosphorylated Spo0A (Spo0A∼P) accumulation leads to the production of SinI, an antagonist of SinR. SinR is a transcriptional repressor that keeps the matrix genes shut off when conditions are not propitious for biofilm growth (29). When environmental cues that induce biofilm formation are present, the kinases are activated, and transcription of the matrix genes is induced via this signal transduction pathway.

We and others have demonstrated the existence of a correlation between B. subtilis root colonization, biofilm formation, and protection against pathogens on tomato plants and in Arabidopsis thaliana (15, 30). However, the expression of the matrix genes was not examined in such biofilms. We recently reported that B. subtilis forms biofilms on tomato roots in a KinD-dependent fashion and that tomato root exudates were able induce B. subtilis biofilm in vitro (31). The tomato exudates contain L-malic acid, which triggers biofilm formation at high concentrations but is unlikely the sole active component of these exudates. Malic acid is also secreted by A. thaliana after infection, but not during regular growth (32); thus, it is not involved in B. subtilis biofilm formation on this plant in the absence of pathogens.

Here, we demonstrate that many genes involved in biofilm formation in vitro are also involved in colonization of A. thaliana roots and that requirement for the kinases is different from for tomato plants. Microscopic imaging showed that matrix genes were expressed in cells colonizing A. thaliana roots and that matrix production was essential for root colonization. Further, we showed that certain plant polysaccharides, usually localized at the surface of the plant cell wall, serve as a cue that stimulates the formation of B. subtilis biofilms. The biofilm matrix was necessary for both root colonization and biofilm formation in vitro in response to plant polysaccharides. In addition, we showed that plant polysaccharides could be metabolized and used as a carbohydrate source to build the EPS component of the B. subtilis matrix. However, monosaccharide constituents of these polysaccharides do not act as potent biofilm inducers. Thus, plant polysaccharides stimulate biofilm formation in B. subtilis in two ways: (i) by inducing matrix gene expression and (ii) by acting as a substrate that is processed and incorporated into biofilm EPS matrix. Finally, plant polysaccharides also robustly stimulate biofilm formation in well-studied plant growth-promoting Bacillus strains.

Results

A. thaliana Root Colonization by B. subtilis Requires Extracellular Matrix.

One of the key requirements for biofilm formation is the secretion of an extracellular matrix that holds the cells together. In incipient B. subtilis biofilms, this matrix is produced by a subpopulation of cells that are genetically identical but phenotypically distinct from the rest of the population (33, 34). The differentiation of these matrix-producing cells can be followed by using a wild-type strain (B. subtilis NCIB3610) expressing a yellow fluorescent protein (yfp) gene under the control of the P_tapA promoter (P_tapA-yfp), which reports expression of the genes encoding the major protein component of the matrix, TasA (34, 35). Using this strain, we analyzed whether colonization of A. thaliana roots by B. subtilis involves induction of the matrix reporter and therefore formation of biofilms akin to those observed in vitro. To perform this experiment, 6-d-old A. thaliana seedlings were added to individual wells of a 48-well plate containing a minimal medium supplemented with a very low concentration (0.05%) of the readily metabolized carbon source glycerol [minimal salts nitrogen glycerol (MSNg) medium] to allow some bacterial growth and thus delay spore formation. Importantly, in this medium, only a few cells express matrix genes (Fig. S2) due to the stochastic switch (36), which is not sufficient to promote biofilm formation in the absence of the plant. Each well was inoculated with B. subtilis harboring the P_tapA-yfp matrix reporter to a very low cell density (final OD₆₀₀ 0.02, ∼10⁷ cfu/mL). This low-density bacterial cell count was important to maintain plant health and to assure that any observed root colonization was the result of specific attachment and bacterial growth at the root and not due to nonspecific cell aggregation resulting from excessive numbers of bacteria in the inoculum. Microscopy imaging was performed to examine the presence of matrix-producing cells at 0, 12, 24 and 48 h after inoculation. For the T₀ time point, inoculation of the bacteria is performed on the microscope slide. At this time point, cells that first come in contact with the root do not express P_tapA-yfp reporter, indicating that matrix gene expression is not required for initial contact (Fig. 1). At 12 h, scattered single cells and some small microcolonies were observed on the roots. After 24 h, 3D clusters of cells, typical of biofilms, had developed on the roots. Sporulation began to occur shortly thereafter. By 48 h, many spores were apparent as they are easily distinguished by their smaller size and phase bright appearance. At 12, 24, and 48 h, a significant proportion of the cells fluoresced due to expression of the P_tapA-yfp reporter (Fig. 1, overlay false-colored green). Cells expressing the matrix reporter were predominantly located in clusters due to the production of extracellular matrix that holds the cells together. However, contrary to what is observed in vitro in biofilm-inducing medium [minimal salts glutamate glycerol (MSgg) medium], cells expressing matrix on the root did not form long chains of cells (37). A human hair was used as a control for nonspecific colonization to a surface under these conditions, and no bacteria attached to it after 24 h incubation (Fig. S3). These observations indicate that A. thaliana root colonization by B. subtilis occurs and involves the activation of matrix genes.

Fig. 1.

B. subtilis cells colonizing A. thaliana roots express matrix genes. Wild-type (3610) cells harboring P_tapA-yfp were coincubated with 6-d-old seedlings of A. thaliana and imaged at various time points postinoculation. Shown are overlays of fluorescence (false-colored green) and transmitted light (gray) images. Pictures are representative of at least ten independent roots. Arrows point toward some of the nonfluorescent cells. (Scale bars: 10 μm.)

As both the EPS and the protein component of the matrix (TasA) are necessary to form robust biofilms in vitro, we examined whether these factors were also required for root colonization. To analyze the ability of different strains to colonize roots, we introduced a constitutively expressed yfp or cyan fluorescent protein (cfp) gene into wild-type B. subtilis and strains containing deletions of genes responsible for EPS production (eps), TasA production (tasA), and an eps tasA double mutant. Using the colonization assay described above, we observed that wild-type strains expressing yfp or cfp constitutively formed robust biofilms on roots (Fig. 2). However, mutant strains lacking either or both components of the matrix were incapable of colonizing the root; only rarely a few single cells were visible on the roots. This result is similar to what was observed for B. subtilis colonization of tomato roots (30).

Fig. 2.

B. subtilis forms biofilms on plant roots in an eps tasA-dependent fashion and requires spo0A and sinI. Wild-type (WT) cells constitutively expressing YFP or CFP and various mutant strains constitutively expressing YFP were coincubated with 6-d-old seedlings of A. thaliana. For the bacterial coinoculation, eps and tasA mutant cells were added at a 1:1 ratio. Colonization of the root was observed after 24 h. Overlays of fluorescence (false colored green for YFP or blue for CFP) and transmitted light images (gray) are shown. Pictures are representative of at least twelve independent roots. (Scale bars: 50 μm.)

Mutants for eps and tasA have been previously shown to complement each other extracellularly (25). This complementation means that biofilms form when the two strains are cocultured. To determine whether the individual eps or tasA mutants were capable of extracellular complementation during root-associated growth, the tasA mutant constitutively expressing cfp was inoculated at a 1:1 ratio with the eps mutant that constitutively expressed yfp. The mutant strains complemented each other and restored wild-type–like colonization; patches of both mutants were observed interspersed on the roots (Fig. 2). These results indicate that both TasA and EPS are required for the formation of a robust biofilm on plants, similar to what has been observed previously in vitro.

As described in the Introduction, a signal transduction pathway—in which the master regulator Spo0A plays a central role—is responsible for activating the transcription of matrix genes in vitro. When the concentration of Spo0A∼P reaches a threshold level, sinI is transcribed (29, 38). The sinI gene product, SinI, derepresses matrix gene transcription by antagonizing the activity of the SinR repressor (Fig. S1). As the matrix is synthesized, biofilm formation ensues. To determine whether this regulatory cascade was involved in biofilm formation during root colonization, spo0A or sinI mutants harboring the constitutive yfp reporter were assessed for their capacity to colonize A. thaliana roots. Neither sinI nor spo0A mutants were able to colonize roots at all (Fig. 2). These results demonstrate the importance of this signal transduction pathway in regulating biofilm formation during root colonization.

Plant Polysaccharides Act as an Environmental Cue for Biofilm Formation.

Because B. subtilis formed biofilms on A. thaliana roots in a medium that does not otherwise induce biofilm formation, we hypothesized that the plant produced a compound that stimulated biofilm formation. This compound could be either secreted or associated with the root. To identify such a compound, we examined the capacity of root exudates and plant extracts to induce B. subtilis biofilm formation. The biofilm-forming activity of exudates and extracts was determined using a standard pellicle assay where cells form a floating biofilm at the air–medium interface. Root exudates were prepared by growing A. thaliana seedlings for 2 wk in 0.2× minimal salts nitrogen (MSN) medium. The medium was then harvested, filter-sterilized, and concentrated; this preparation was termed “root exudates.” The same plants used for exudate preparation were then crushed after freezing in liquid nitrogen. The resulting powder was resuspended in water, filter-sterilized, and concentrated; this solution was termed “plant extracts.” The capacity of the plant extracts and/or the root exudates to influence biofilm formation by B. subtilis was investigated in MSN medium supplemented with 0.5% cellobiose (MSNc). Cellobiose is a product of partial cellulose hydrolysis similar to what could be found on plants and was used as a carbon source to support growth during the assay. The final exudate or extract used for the biofilm assay was concentrated fivefold as bacteria close to the roots are likely to encounter significant amounts of such materials. In the wells containing no addition (−) or root exudates, cells grew well but did not form biofilms as indicated by the absence of a pellicle (Fig. 3). This growth is easily visualized compared with the control “no cells,” which contains only medium. In contrast, plant extracts were able to induce biofilm formation as evidenced by the formation of a biofilm at the air/liquid interface (Fig. 3). The pellicle gives a more opaque appearance to the culture than planktonic growth. This result confirms that a substance(s) that triggers B. subtilis biofilm formation is present on or in plants.

Fig. 3.

Plant extracts but not root exudates induce biofilm formation. Pellicle formation of wild-type (3610) cells in MSNc with no addition (−) or with root exudates or plant extracts added at the onset of the assay. Images are top-down view of wells and were taken after 24 h at 30 °C. The “No cells” control is shown to highlight the appearance of bacterial cell growth in the inoculated wells.

Some of the first molecules that B. subtilis encounters on the plant root are the polysaccharides from the plant cell wall. Therefore, we hypothesized that plant polysaccharides could be some of the components in plant extracts that trigger biofilm formation by B. subtilis. We tested this hypothesis with a pellicle assay in which MSNc was supplemented with various purified plant polysaccharides (arabinogalactan, pectin, or xylan) added at a final concentration of 0.5%. All three of the plant polysaccharides tested were capable of inducing pellicle formation after 24 h, as displayed in Fig. 4A. To quantify pellicle formation, we developed a pellicle weight assay using preweighed wells with a mesh bottom. These wells were inserted in 24-well plates where the assay was performed, and, after 24 h, the pellicle was separated from planktonic growth by pulling out the mesh-bottomed well. The pellicle was dried on the mesh, and total weight was measured. We observed that the final mass of material present on the mesh in medium containing plant polysaccharides was more than twice that of medium with no plant polysaccharides (−) (Fig. 4B). The results of this assay corresponded nicely with the qualitative difference observed in the pellicle images and show that robust pellicle formation occurs in the presence of plant polysaccharides This effect was limited to plant polysaccharides. We tested other (nonplant) polysaccharides, such as dextran and laminarin, and they did not induce pellicle formation (Fig. 4 A and B). A difference between plant polysaccharides and other polysaccharides was also observed when it came to the ability of B. subtilis to use them as sole source of carbon. Indeed, B. subtilis could grow with plant polysaccharides as sole carbon source, but it could not similarly use dextran or laminarin (Fig. S4).

Fig. 4.

Plant polysaccharides promote biofilm formation. (A) Pellicle formation of wild-type cells in MSNc. The indicated di- ,tri-, or polysaccharides were added at a final concentration of 0.5% at the onset of the assay. AG, Arabinogalactan. Images are top-down views of wells and were taken after 24 h at 30 °C. (B) Pellicle weight assay of wild-type cells in MSNc. The indicated di-, tri-, or polysaccharides were added at a final concentration of 0.5% at the onset of the assay. Analysis of variance revealed a significant main group effect between the conditions used [F(10, 75), P < 0.001]. Tukey's post hoc test revealed that arabinogalactan, pectin, and xylan (marked with asterisks) showed greater pellicle mean mass compared with all other conditions. (C) Flow cytometry analysis of wild-type cells harboring the P_tapA-yfp reporter grown in MSNc with indicated additions or no supplements (−). Fluorescence intensity in arbitrary units is shown on the x axis and number of cells is shown on the y axis. Results shown are representative of three independent experiments.

The induction of biofilm formation by plant polysaccharides described above was observed in MSNc medium. In this medium, B. subtilis grows well, and growth was not limited in any of the conditions that did not result in pellicle formation. Nevertheless, to rule out the possibility that the effect of plant polysaccharides could be due to an increased amount of available carbon source, controls with the trisaccharide raffinose and with glycerol were performed. Neither induced biofilm formation (Fig. 4 A and B). Similarly, galactose, arabinose, fucose, xylose, and glucuronic acid added at a 0.5% concentration did not induce pellicle formation (Fig. S5), which further demonstrates that the effect of plant polysaccharides as an environmental cue inducing biofilm formation is not due to increased growth attributed to the presence of additional sugars. Addition of galacturonic acid gave rise to a thin pellicle (Fig. S5), but, because 0.5% pure galacturonic acid has a weaker effect than 0.5% arabinogalactan or pectin, it is likely that the contribution of this monosaccharide to the global effect of arabinogalactan and pectin is minimal. Lactose, which contains galactose residues that are common in arabinogalactans and pectin, and PEG-300, which acts as a control for increased osmolarity (a physical change that occurs upon addition of plant polysaccharides), also had no influence on pellicle formation (Fig. 4 A and B). Thus, the induction of biofilm formation in MSNc medium appears to be a specific response to the addition of plant polysaccharides. Notably, the addition of lower concentrations of plant polysaccharides was incapable of inducing robust biofilm formation; the addition of 0.1% had almost no effect, and we were unable to observe any pellicle when we decreased the concentration to 0.05% (Fig. S6).

Expression of matrix genes in a subpopulation of cells is a hallmark of B. subtilis biofilm formation. To determine whether there was increased expression of matrix genes concomitant with the observed pellicle formation in response to plant polysaccharide, assays were performed using wild-type cells harboring the P_tapA-yfp matrix reporter. After 24 h, all of the cells in a given well were harvested, fixed, and analyzed by flow cytometry. As shown in Fig. 4C, when cells were incubated in the presence of arabinogalactan, pectin, and xylan, a large fraction of the population expressed yfp. All of the other conditions tested, including nonplant polysaccharides, di- and trisaccharides, and osmotic controls, had ample cell growth yet little expression of the P_tapA-yfp reporter. Induction of matrix gene transcription corresponded with the molecules' ability to induce pellicle formation (compare Fig. 4 B and C). Thus, we conclude that plant polysaccharides induce transcription of matrix genes and that this induction leads to biofilm formation.

The specificity of the plant polysaccharides suggests that they can be an environmental cue for biofilm formation. Other known signals for biofilm formation are sensed through histidine kinases that phosphorylate Spo0A, and Spo0A∼P leads to SinI accumulation and matrix gene expression. Thus, we tested whether Spo0A and SinI were required for the bacteria to respond to plant polysaccharides. Deletion of either spo0A or sinI completely abolished plant polysaccharide-induced biofilm formation (Fig. 5A). These results are consistent with the observed defect in root colonization of these two mutants (Fig. 2). Importantly, whereas there was no biofilm formation by the spo0A or sinI mutants, cells were still able to grow dispersed under these conditions. This growth was obvious when comparing the wells inoculated with the mutants with uninoculated control wells that contain the polysaccharides (Fig. 5A). Xylan in solution is milky, as observed in the uninoculated control, and there is colloidal pectin showing some debris. Quantification of pellicle weight further supports the observation that sinI and spo0A mutants are unable to form biofilm in the presence of plant polysaccharides (Fig. 5B).

Fig. 5.

The master regulator Spo0A and the SinI antirepressor are involved in responding to plant polysaccharides. (A) Top-down view of pellicle assay in which the indicated regulatory mutant cells were incubated for 24 h at 30 °C in the presence of arabinogalactan (AG), pectin, or xylan in MSNc medium. “No cells” controls show that mutant cells are able to grow but do not form a pellicle. (B) Weight quantification of pellicles formed in the same conditions as in A. A two-way analysis of variance revealed a significant difference between the mutants and wild type [F(2, 27), P < 0.001]. Tukey's post hoc test revealed that sinI and spo0A (marked with asterisks) showed lower pellicle mean mass compared with wild type in the same conditions.

We next asked whether any of the five kinases (KinA–KinE) known to phosphorylate Spo0A was responsible for sensing plant polysaccharides. The pellicle formation assay demonstrated that all of the single kinase deletions were still capable of forming pellicles in response to these three plant polysaccharides (Fig. 6 A and B). However, pellicles produced by kinC and kinD mutants appeared more fragile to the naked eye, and concomitantly a kinCD double mutant was defective in forming pellicles in response to arabinogalactan and pectin, an effect that could be quantified using the pellicle weight assay (Fig. 6 A and B). These results point toward an involvement of KinC and KinD in arabinogalactan- and pectin-mediated biofilm formation. Interestingly, pellicles formed in xylan were not affected by the deletion of any of the kinase genes, suggesting that xylan acts through an as yet unidentified pathway. Given the fact that some plant polysaccharides are sensed by KinC and KinD, we wondered whether these genes were important for plant colonization. Thus, we tested the single kinase mutants as well as the kinCD double mutant for their ability to colonize A. thaliana roots. All of the single kinase deletion mutants were able to colonize the root as efficiently as the wild type (Fig. S7). The kinCD double mutant appeared only marginally defective by visualization, and this observation was confirmed by averaging fluorescence of 70 pictures from 12 roots. This result indicates that there are likely redundant plant molecules acting as environmental cues to trigger biofilm formation on plant roots (Fig. S7).

Fig. 6.

KinC and KinD are involved in sensing and responding to plant polysaccharides. (A) Top-down view of pellicle assay in which the indicated regulatory mutant cells were incubated for 24 h at 30 °C in the presence of arabinogalactan (AG), pectin, or xylan in MSNc medium. (B) Weight quantification of pellicles formed in the same conditions as in A. A two-way analysis of variance revealed a significant difference between the mutants marked with an asterisk and wild type [F(6, 63), P < 0.001]. Tukey's post hoc test revealed that kinCD in the presence of AG and pectin (marked with asterisks) showed lower pellicle mean mass compared with wild type in the same conditions, but that was not the case in the presence of xylan.

Plant Polysaccharides Are Used as a Carbon Source for the Synthesis of Matrix EPS.

Using the Carbohydrate Active Enzymes database (www.cazy.org/), we carried out a bioinformatic search to identify glycosyl hydrolases that might be encoded in the B. subtilis genome (using strain 168 as a reference) (39). The search retrieved over 40 different predicted glycosyl hydrolases that could digest plant polysaccharides and thus allow them to be used as a carbon source to sustain cell growth (as was observed in Fig. S4). We therefore tested whether plant polysaccharides could also be used as a carbohydrate source for building the EPS component of the biofilm matrix. We recently demonstrated that production of the matrix EPS requires the synthesis of UDP-galactose, which likely serves as a substrate to be incorporated into the EPS (40). UDP-galactose can be generated via two different pathways. The first involves the production of UDP-glucose and its subsequent conversion into UDP-galactose by the enzyme GalE. The second pathway necessitates direct modification of galactose into UDP-galactose by GalK and GalT (Fig. 7A). In a defined medium where glycerol and glutamate are the only carbon sources (MSgg; see Materials and Methods), a galKT deletion mutant, but not a galE mutant, is capable of forming pellicles (Fig. S8). Under these conditions, the galE defect can be rescued if an external source of galactose is provided (ref. 40 and Fig. S8). Because many plant polysaccharides contain galactose residues, it seemed possible that they could be incorporated into the matrix EPS via the GalK and GalT pathway. To test this, galE and galKT mutants were assayed for their capacity to form biofilms in the presence of plant polysaccharides. This assay was performed in the MSNc medium, which does not induce biofilm formation and lacks galactose, but contains cellobiose as a carbon source that permits growth. As shown in Fig. 7B, a galE mutant in the presence of arabinogalactan formed a pellicle similar to that formed by the wild-type strain, suggesting that the galactose residues contained in this plant polysaccharide could have been incorporated in the matrix EPS via the galKT pathway. The same observation applied for pectin, which led to the formation of a robust pellicle by both the wild type and the galE mutant (Fig. 7B). The pellicle weight assay further demonstrated no statistical difference between wild type and the galE mutant in the presence of arabinogalactan and pectin, whereas the galE galKT mutant formed much weaker pellicles. These observations are consistent with the fact that both of these polysaccharides are known to contain many galactose residues (41). Xylan is a polymer of xylose residues that contains few galactose residues. Accordingly, the visual appearance of the galE mutant showed an incomplete pellicle in the presence of xylan, and this decreased pellicle was corroborated by the quantification of its mass that is statistically different from the mass of a wild-type pellicle in the presence of xylan. The galKT mutant was able to form pellicles in the presence of all of the plant polysaccharides, likely due to the utilization of sugar residues other than galactose as a source for UDP-glucose that could be converted to UDP-galactose by GalE (Fig. 7 B and C). Finally, as expected, a strain combining both galE and galKT deletions was defective in biofilm formation regardless of the addition of plant polysaccharides (Fig. 7 B and C). Thus, in addition to inducing matrix gene expression, plant polysaccharides can also serve as a source of carbon to build the matrix EPS. To examine whether B. subtilis was also able to obtain galactose from the plant, colonization assays were performed with galE, galKT, and galE galKT mutants. Interestingly, although they did not colonize to wild-type levels, galE cells were still able to form microcolonies on plant roots (Fig. 7D). As expected, the galE galKT mutant was completely defective in root colonization. The addition of 0.05% galactose in the culture medium restored root colonization to a galE mutant but had no effect on galE galKT colonization (Fig. 7D). These results demonstrate that Arabidopsis roots can provide enough galactose to B. subtilis to sustain limited root colonization.

Fig. 7.

Plant polysaccharides provide a substrate that is incorporated into the matrix EPS. (A) A proposed model for the metabolism of galactose that is incorporated into the matrix EPS. GalK and GalT convert exogenous galactose to UDP-galactose, which is used in EPS production. GalE converts UDP-glucose into UDP-galactose, producing UDP-galactose from central metabolism. (B) Top-down view of pellicle assay in which the indicated mutants were incubated for 24 h at 30 °C in the presence of arabinogalactan (AG), pectin, or xylan in MSNc medium. Results are representative of three experiments. (C) Weight quantification of pellicles formed in the same conditions as in B. A two-way analysis of variance revealed a significant difference between the mutants [F(6, 63), P < 0.001]. Tukey's post hoc test revealed that in the presence of AG and pectin, only galE galKT mutant (marked with asterisks) showed lower pellicle mean masses compared with wild type (WT), but in presence of xylan both galE and galE galKT are significantly different from WT. (D) Root colonization of 3610, galE, galKT, and galE galKT mutants constitutively expressing YFP. When indicated, galactose is added at a concentration of 0.05%. Pictures are representative of at least 16 independent roots for those without galactose and 10 for those with galactose. (Scale bars: 50 μm.)

A. thaliana Arabinogalactan Proteins Are Potent Inducers of B. subtilis Biofilm Formation.

The various plant polysaccharides used throughout this study are purified from different plants, yet all of them induced pellicle formation. However, our colonization experiments were performed on A. thaliana, and thus we wanted to confirm that polysaccharides from this model plant could trigger B. subtilis biofilm formation. As shown in Fig. 8, arabinogalactan proteins isolated from A. thaliana trigger the formation of a robust pellicle. The induction effect was robust and occurred at a concentration of 0.05%, which was a concentration in which commercially available arabinogalactan was not active. These results demonstrate that arabinogalactan proteins present in the A. thaliana cell wall are potent inducers of B. subtilis biofilm formation and thus may be able to induce the expression of matrix genes when the bacteria are in contact with A. thaliana roots.

Fig. 8.

A. thaliana arabinogalactan proteins induce biofilm formation. (A) Pellicle formation of wild-type cells in MSNc in the presence of 0.05% or 0.5% commercial arabinogalactan (AG) or purified A. thaliana arabinogalactan proteins (A. thaliana AGPs). Images are top-down views of wells and were taken after 24 h at 30 °C. (B) Weight quantification of pellicles formed in the same conditions as in A. A two-way analysis of variance revealed a significant difference between the conditions marked with an asterisk and the untreated sample [F(4, 15), P < 0.001]. Tukey's post hoc test revealed 0.5% AG, and both concentrations of A. thaliana AGPs showed higher pellicle mean mass compared with the untreated well.

Plant Polysaccharides Influence Biofilm Formation in Various Plant Growth-Promoting Bacillus Species.

Many strains of B. subtilis and closely related species are known to have beneficial effects on plant growth (18). We thus wanted to test whether plant polysaccharides would also act as an environmental cue for biofilm formation in such strains. Using the same pellicle formation assay as described earlier, B. subtilis GB03 and Bacillus amyloliquefaciens FZB42 responded robustly to plant polysaccharides, producing heavily wrinkled pellicles in the presence of arabinogalactan, pectin, and xylan (Fig. 9). These results show that the ability to recognize plant polysaccharides as an environmental cue for biofilm formation is conserved in plant growth-promoting B. subtilis/amyloliquefaciens strains.

Fig. 9.

Biofilm formation by plant growth-promoting Bacillus strains is influenced by plant polysaccharides. Pellicle formation by B. subtilis 3610, the plant growth-promoting strain B. subtilis GB03, and B. amyloliquefaciens FZB42 in MSNc with arabinogalactan (AG), pectin, and xylan. Top-down view of pellicle after incubation for 24 h at 30 °C. Results are representative of three experiments.

Discussion

We have shown that colonization of A. thaliana roots by B. subtilis displays striking similarity to the process of biofilm formation in vitro. A large subpopulation of cells adhering to A. thaliana roots expressed genes whose products produce an extracellular matrix. The production of both the EPS and protein components of the extracellular matrix is crucial for plant colonization as cells defective in either component could not form biofilms on the root. In addition, we found that plant extracts stimulated biofilm formation and matrix gene expression in vitro. Further, we showed that purified plant polysaccharides, namely arabinogalactan, pectin, and xylan, but not their subunits or other complex polysaccharides of nonplant origin, act as an environmental cue to induce matrix gene expression, which in turn leads to biofilm formation. These plant polysaccharides can also be digested, converted to UDP-galactose, and incorporated into the matrix EPS. Thus, plant polysaccharides act both as an environmental cue and as a substrate for B. subtilis matrix synthesis.

The eps and tasA mutants almost completely failed to colonize the root, suggesting that both matrix components are critical for root colonization. EPSs were shown to be important for root colonization in many bacterial species, such as for Azospirillum brasilense, Gluconacetobacter diazotrophicus, Herbaspirillum seropedicae, Agrobacterium tumefaciens, Sinorhizobium meliloti and Pseudomonas fluorescens (42–48). In a Gluconacetobacter mutant in which the first step of EPS biosynthesis was inactivated, exogenous addition of EPSs could partially restore biofilm formation on the roots (45). Similarly, when we mixed B. subtilis eps and tasA mutants, which were individually incapable of plant colonization, we observed the formation of robust biofilms containing both strains on plant roots. This result suggests that both the EPS and protein components of the B. subtilis matrix can be provided extracellularly to support root colonization. The B. subtilis TasA protein is not the first example of a bacterial protein that is involved in plant colonization. In Pseudomonas putida, the matrix large adhesion protein F (LapF) is important for colonization, and a LapF mutant has reduced competitive fitness on alfalfa roots compared with a wild-type strain (49).

We have identified the plant polysaccharides pectin, arabinogalactan, and xylan as an environmental cue triggering biofilm formation by plant growth-promoting B. subtilis. This strong effect on biofilm formation was specific to these three plant cell wall polysaccharides because xyloglucan had a weaker capacity to induce pellicle formation (Fig. S9). We also tested the soluble derivative of cellulose carboxymethylcellulose, and, although it could not induce biofilm formation (Fig. S9), we cannot distinguish whether this lack of effect is due to the artificial modification of cellulose or because cellulose itself has no effect.

Interestingly, in B. subtilis, whereas the effect of arabinogalactan, pectin, and xylan was to promote biofilm formation, the mechanisms by which this effect occurs appear to be different depending on the polysaccharide. Statistical analysis of the masses of wild-type pellicles showed that xylan promoted thicker biofilms whereas there was no difference between the effect of arabinogalactan and pectin. This result is consistent with the fact that these polysaccharides have very different structures. Xylan is a polymer of xylose, a pentose, to which side chains are added in A. thaliana. It is present mostly in fiber cells, contrary to pectin and arabinogalactans, which are generally found throughout the plant and thus are more likely to be present on the roots (41, 50). Consequently, we propose that, for surface colonization of A. thaliana, pectin and arabinogalactans may be the major players. However, B. subtilis is very often found growing associated with decaying plant material (for review, see ref. 51), in which case xylan would more likely be exposed. It is possible that xylan serves as a signal for colonization and biofilm formation on decaying plants, which also serve as a rich source of carbon.

Pectin is a complex polysaccharide composed mainly of galacturonic acid, branched arabinan, and 1,4-β-galactans and constitutes about 40% of the A. thaliana primary cell wall (41, 50). Finally, arabinogalactans are attached to the arabinogalactan proteins (AGPs), which play important roles in development of the plant and its interaction with microorganisms (52). Notably, Vicré et al. showed that colonization of A. thaliana root border cells by Rhizobium sp. YAS34 is disrupted by treating the roots with the β-glucosyl Yariv reagent, which binds and precipitates arabinogalactans (53). Similarly, an A. thaliana mutant for the Arabinogalactan-protein AtAGP17 is resistant to A. tumefaciens transformation, and further investigation of that phenomenon showed that arabinogalactan moieties are involved in the first attachment step of Agrobacterium (54). These observations suggest a crucial role for arabinogalactan in mediating the attachment of various rhizobacteria to the root.

We discovered that AGPs purified from A. thaliana were able to trigger biofilm formation (Fig. 8). This result suggested that AGPs could be a factor in Arabidopsis plant extracts that can stimulate biofilm formation. Are the AGPs, pectins and xylan serving to induce biofilm formation when B. subtilis encounters the plant? At present, this question is not readily answered. One possibility might be to modify the roots chemically to disrupt the AGPs using the β-glucosyl Yariv reagent (53). However, such treatment would leave the pectin still present, and thus biofilms might still form. Genetic approaches to modifying the root cell wall polysaccharides could also be considered (41, 50). Although there are mutants that affect cell wall polysaccharides, e.g., rat4, which affects expression of the cellulose synthase-like CSLA9 (55), at present we are unaware of any Arabidopsis mutants that completely eliminate AGPs, pectin, and xlyan. A more direct test would be to obtain B. subtilis mutants unable to sense plant polysaccharides in vitro and then test them in planta. Such mutants do not currently exist although they would be a very useful tool for this system.

Although bacterial EPSs have been known to have many functions in signaling to plants (56, 57), this report presents previously undescribed evidence of plant polysaccharides acting as an environmental cue for bacterial biofilm formation. However, it was reported in Clostridium thermocellum that plant polysaccharides could activate a transcriptional response via a set of seven sigma (SigI) and anti-sigma (RsgI) factors (58). Each RsgI homolog contains a carbohydrate-binding module that senses specific plant polysaccharides and induces a conformational change on the intracellular anti-σ domain, resulting in the release of the alternative σ factor and transcriptional activation of its genes (59). B. subtilis possesses one sigI-rsgI homolog; however, the encoded RsgI protein does not possess the carbohydrate-binding module. Also, deletion of the operon encoding both proteins did not have an effect on biofilm formation with plant polysaccharides (Fig. S10).

Although biofilm matrix components from several bacterial species have been implicated in root colonization, before this work, regulation of biofilm formation had not been shown to have a direct impact on A. thaliana root colonization. Here, we have shown that the same signal transduction pathway is used in vitro and in vivo to regulate biofilm formation. Indeed, we found that the transcriptional regulator Spo0A and the matrix anti-repressor SinI were both required for root colonization. SinI has also been recently shown to be required for tomato roots colonization (30). Moreover, both SinI and Spo0A are essential for the biofilm-inducing effect of plant polysaccharides, suggesting that plant polysaccharides act through the Spo0A/SinI signal transduction pathway. The sensors in this pathway are five histidine kinases (28). Although the kinCD double mutant formed much weaker pellicle in the presence of arabinogalactan and pectin, these mutations had little to no impact on biofilm formation in the presence of xylan. Thus, there may be an alternative pathway involved in sensing xylan. This alternative pathway may also be at play on A. thaliana plant roots where the kinCD mutant is only marginally defective in colonization. Interestingly, KinC responds to potassium leakage that is induced by the self-produced molecule surfactin (60). In addition, it has been previously shown that a B. subtilis mutant unable to make surfactin is defective in root colonization (15). The discrepancy between the kinC and the surfactin mutant in root colonization indicates that the main role of surfactin on the root is not likely to be its capacity to signal through KinC.

Recently, we have also investigated B. subtilis biofilm formation on tomato roots (31). In both tomato and Arabidopsis, biofilm formation on the root depended on matrix genes and the Spo0A/SinI regulatory pathway. However, the kinase requirement for such colonization differed from one plant to another. Whereas, with A. thaliana roots, the kinCD deletion had only a moderate effect on biofilm formation, the same deletion completely abrogated colonization of tomato roots. Thus, it appears that B. subtilis biofilm formation on roots involves different kinases and possibly different environmental cues depending on the plant species. Such a result is not unprecedented; Bacillus cereus colonization also varies according to the strain of tomato plant that is assayed (61).

In addition to acting as an environmental cue for biofilm formation, plant polysaccharides can be digested and their products used as building blocks for the formation of the EPS portion of the B. subtilis biofilm matrix. Indeed, B. subtilis possesses a large number and variety of secreted enzymes that are able to degrade plant polysaccharides such as xylan and arabinogalactans and to import mono-, di-, and trisaccharides into the cell (62–64). As the synthesis of matrix polysaccharide requires abundant carbon input, the fact that B. subtilis can use galactose from the plant polysaccharides as a building block for its matrix reflects admirable energy conservation.

The biofilm-enhancing effect of plant polysaccharides is not limited to B. subtilis strain NCIB 3610, which we used throughout most of the studies presented here. We used this strain because most of our prior work on biofilm formation was done with it and because of its genetic manipulability. However, we did observe that the plant growth-promoting strains B. subtilis GB03 and B. amyloliquefaciens FZB42 produced more robust biofilms in the presence of plant polysaccharides. Importantly, Bacillus amyloliquefaciens FZB42 harbors genes encoding for arabinogalactan hydrolysis, a sugar importer and galE, galK and galT. All of these genes are clustered in what appears to be a single operon (65). This observation further strengthens the idea that the ability to sense plant polysaccharides to trigger matrix gene expression and to use them as substrate to build the matrix is conserved among B. subtilis and closely related species. Indeed, the capacity to form biofilms when in contact with plant polysaccharides could be an advantageous trait for plant growth-promoting bacteria, serving to enhance colonization of the roots. Accordingly, many Bacillus species isolated from plant roots were able to form robust biofilms in the appropriate medium (66). Further comparison of GB03, FZB42, and NCIB 3610 genomes will provide clues to the genetic requirements for this trait. Although the beneficial effects on plants by Bacillus species have been largely demonstrated, the molecular details regulating the lifestyle of these bacteria when in contact with the root are still not well understood. The role of plant polysaccharides on bacterial biofilm formation provides one of the first clues as to how the plant might stimulate its colonization by this beneficial microorganism.

Materials and Methods

Strains, Media and Culture Conditions.

Strains used in the study are listed in Table S1. The B. subtilis strain NCIB 3610 was used as the wild-type strain because the common B. subtilis 168 laboratory strain contains mutations that impair its ability to form biofilm (67). Other Bacillus species were acquired as kind gifts from Joseph W. Kloepper (Auburn University, Auburn, AL; GB03 strain) and Michael Fischbach (University of California, San Francisco, CA; strain FZB42), and through the Bacillus Genetic Stock Center. For routine growth, cells were propagated on Luria-Bertani medium (LB). When necessary, antibiotics were used at the following concentrations: MLS (1 μg⋅mL⁻¹ erythromycin, 25 μg⋅mL⁻¹ lincomycin), spectinomycin (100 μg⋅mL⁻¹), tetracycline (10 μg⋅mL⁻¹), chloramphenicol (5 μg⋅mL⁻¹), and kanamycin (10 μg⋅mL⁻¹). Purified plant polysaccharides were from Sigma except the A. thaliana arabinogalactan proteins, which were a generous gift of Eva Knoch (Laboratory of Naomi Geshi, Department of Plant and Environmental Sciences, University of Copenhagen, Copenhagen, Denmark), purified according to Tryfona et al. (68). Plant polysaccharides were used at a 0.5% (wt/vol) final concentration except if indicated otherwise.

For pellicle assays, cells were cultured from 1-d-old colonies resuspended in 3 mL of LB at 37 °C. After 2 h, cells were diluted 1:100 in 3 mL of LB, and this dilution was repeated at least one more time. After the last dilution, cells were harvested at OD₆₀₀ < 0.5 and adjusted to a final OD₆₀₀ of 0.3. Pellicle assays were performed in 1 mL of medium in a 24-well plate, to which 13.5 μL of cells were added. For root exudates and plant extracts, 300 μL in 48-well plates were used, and 4 μL of cells were added. Plates were incubated at 30 °C for 24 h or 48 h. Medium used throughout this study is MSN (5mM potassium phosphate buffer pH7, 0.1M Mops pH7, 2mM MgCl₂, 0.05mM MnCl₂, 1μM ZnCl₂, 2 μM thiamine, 700 μM CaCl₂, 0.2% NH₄Cl) supplemented with either 0.05% glycerol (MSNg, for root colonization) or 0.5% cellobiose (MSNc, for pellicle assays). These media are a variation of the biofilm-inducing MSgg medium, which contains additional components that induce biofilm formation (37). Pellicle assays with the various bacillus strains were performed in 24-well plates with 1 mL of MSNc medium supplemented or not with the plant polysaccharides.

A. thaliana used in this study is of the Col-0 ecotype (a kind gift of Fred Ausubel, Harvard Medical School, Boston, MA). Seeds were surface-sterilized with 70% (vol/vol) ethanol followed by 0.3% sodium hypochlorite (vol/vol) and germinated on Murashige-Skoog medium (Sigma) 0.5% agar with 0.05% glucose in a growth chamber at 25 °C.

Strain and Plasmid Construction.

The long-flanking homology PCR technique was used for creating deletion mutations. Primers used for the kinE::cm gene replacement mutant were as follows (5′-3′): HV102-CAAGGAACATCGGTAAGAATAC, HV103- CTTGATAATAAGGGTAACTATTGCCCAGCTCCGAGTTTGTCTGG, HV104- GGGTAACTAGCCTCGCCGGTCCACGGTTTTCCATATTACGCTTCCTG, and HV105- CTCATTTGGAGCCGAGTCAG. The constitutively fluorescent (YFP) strain PB133 is derived from P_hyperspank promoter (a kind gift of Edgardo Sanabria-Valentín, Harvard Medical School, Boston, MA). The constitutively fluorescent (CFP) strain HV1142 was constructed by inserting integration vector pHV119 into the amyE gene on the chromosome. pHV119 was made by amplifying P_spac from plasmid pDGIEF (69) using primers (5′-3′): HV43b-CCGGAATTCTACACAGCCCAGTCC and HV44b-GGCAAGCTTAATTGTTATCCGCTCAC. P_spac was then cloned upstream of the cfp gene in the amyE integration plasmid pKM008 using EcoRI and HindIII. The LacI repressor gene was not included in the plasmid, making CFP fluorescence constitutive. Plasmids with promoter fusion or PCR products for gene deletions were transferred to B. subtilis strain 168 by natural competence (70). Transformants were selected with the appropriate antibiotics for a double crossover recombination at the amyE or the lacA locus (71) (72). Promoter fusions or gene deletion were then transferred to the strain NCIB3610 or other appropriate mutant strain by SPP1-mediated generalized transduction (73).

Root Colonization Assay.

Six-day-old seedlings were transferred to 300 μL of MSNg in a 48-well plate. Medium hosting the plant was then inoculated at OD₆₀₀ = 0.02 with B. subtilis grown for 3 h, and put on an orbital shaker at 100 rpm in the greenhouse for 24 h.

Image Capture and Microscopy.

To view bacteria on the root surface, seedlings were examined with a Nikon Eclipse TE2000-U microscope equipped with a 20× Plan Apo objective (Figs. 2 and 3) or 60× Plan Apo oil objective (Fig. 1), and pictures were taken with a Hamamatsu digital camera model ORCA-ER. Fluorescence signal was detected using a CFP/YFP dual-band filter set (Chroma #52017). All images were taken at the same exposure time, processed identically for compared image sets, and prepared for presentation using MetaMorph and Photoshop Software. Each image is representative of at least 12 root colonization assays performed in three independent experiments; occasionally there was variation for a given sample. Pellicles were photographed with a 0.8× objective using a Zeiss Stemi SV6 Stereoscope connected to a color AxioCam.

Root Exudates and Plant Extracts Preparation.

To collect root exudates, 10-d-old seedlings were transferred to a 12-well plate containing 3 mL of 0.2× MSNg, four seedlings per well, and incubated for 2 wk in the greenhouse on an orbital shaker at 100 rpm. Plants were then removed, and the contents of five wells were pooled and filtered with a 0.45-μM filter and concentrated 10 times using a speedvac. For plant extracts, the 20 plants used to make the exudates were frozen with liquid nitrogen and ground with mortar and pestle, resuspended in 4 mL of water, filtered with a 0.45-μM filter, and concentrated 10× using a speedvac. Extracts and exudates were used at a 5× final concentration for the pellicle assay.

Flow Cytometry.

For flow cytometry analysis, cells were grown as pellicles. Pellicles were broken and cells harvested by repeated pipetting up and down. Subsequent steps are as described previously (34).

Pellicle Weight Assay.

For this assay, preweighed PELCO prep-eze individual well for 24-well plates with a mesh bottom (opening size 420 μM) (Ted Pella) were put in the wells of multiwell dishes. Media and cells were added subsequently, and pellicles were allowed to develop for 24 h at 30 °C. Individual wells were then removed, dried, and weighed. Each figure is an experiment performed in at least a quadruplicate, and, whereas the absolute values varied between experiments, the trends were always the same.

Statistical Analyses.

Statistical analyses were performed using PRISM 6 software (Graphpad). Comparisons were done using one-way analysis of variance (ANOVA), followed by Tukey’s multiple comparison test (set at 5%).