Special Issue Meeting Presentation

The Two Chemotaxis Clusters in Caulobacter crescentus Play Different Roles in Chemotaxis and Biofilm Regulation

Cécile Berne, Yves V. Brun
George O’Toole, Editor
DOI: 10.1128/JB.00071-19

ABSTRACT

The holdfast polysaccharide adhesin is crucial for irreversible cell adhesion and biofilm formation in Caulobacter crescentus. Holdfast production is tightly controlled via developmental regulators, as well as via environmental and physical signals. Here, we identify a novel mode of regulation of holdfast synthesis that involves chemotaxis proteins. We characterized the two identified chemotaxis clusters of C. crescentus and showed that only the previously characterized major cluster is involved in the chemotactic response toward different carbon sources. However, both chemotaxis clusters encoded in the C. crescentus genome play a role in biofilm formation and holdfast production by regulating the expression of hfiA, the gene encoding the holdfast inhibitor HfiA. We show that CheA and CheB proteins act in an antagonistic manner, as follows: while the two CheA proteins negatively regulate hfiA expression, the CheB proteins are positive regulators, thus providing a modulation of holdfast synthesis and surface attachment.

IMPORTANCE Chemosensory systems constitute major signal transduction pathways in bacteria. These systems are involved in chemotaxis and other cell responses to environment conditions, such as the production of adhesins to enable irreversible adhesion to a surface and surface colonization. The C. crescentus genome encodes two complete chemotaxis clusters. Here, we characterized the second novel chemotaxis-like cluster. While only the major chemotaxis cluster is involved in chemotaxis, both chemotaxis systems modulate C. crescentus adhesion by controlling expression of the holdfast synthesis inhibitor HfiA. Here, we identify a new level in holdfast regulation, providing new insights into the control of adhesin production that leads to the formation of biofilms in response to the environment.

INTRODUCTION

In their natural habitat, most bacteria are organized in complex surface-associated multicellular communities known as biofilms. The first step of biofilm formation is the reversible adhesion of a few single cells to a surface. When conditions are favorable, these attached cells produce adhesin molecules, which strengthen the interaction with the surface. The cells then divide to form multicellular microcolonies, which eventually develop into a mature biofilm (1). Communal life on a surface is believed to be beneficial, as it provides protection from predators and xenobiotic stresses (2). The environment at the surface is highly heterogenous, with the presence of various compounds adsorbed on the surface and the formation of gradients near it (1). To initiate attachment, bacteria must approach the surface either by passive transport or by active swimming (1). Both active swimming toward the surface and initial surface attachment can be biased by environmental cues and chemotaxis (3, 4). For example, chemotaxis is involved in the colonization of biotic (5, 6) and abiotic (7–10) surfaces and in cell-cell aggregation (11, 12). Finally, chemotaxis is also involved in later stages of biofilm formation, as is the case for single Pseudomonas aeruginosa cells that can actively respond to a chemical gradient and subsequently reposition themselves on the surface within a mature biofilm (13).

The central chemotaxis system is composed of a chemoreceptor, or methyl-accepting chemotaxis protein (MCP), and CheW, CheA, and CheY proteins (14–18). These proteins form large complexes of hexagonally packed arrays, localized at the cell pole, with MCPs being transmembrane proteins, and with CheW, CheA, and CheY being located in the cytoplasm (19, 20). The chemotactic signal is sensed by the MCP and transduced to the sensory histidine kinase CheA via the scaffolding protein CheW (Fig. 1A). If the signal is an attractant, CheA autophosphorylation is inhibited, while a repellant signal activates it. Phosphorylated CheA (CheA∼P) serves as a phosphodonor to the CheY and CheB response regulators. CheY∼P interacts with the flagellum apparatus, modulating flagellum rotation and swimming behavior. CheB∼P removes a methyl group from the MCP, reducing the overall activity of the signal transduction cascade and modulating the chemotactic response. Optional accessory proteins have also been linked to the chemotaxis apparatus and play auxiliary roles in chemotaxis regulation, such as the phosphatases CheC, its homolog CheX, and CheZ (CheY∼P hydrolyzation), the glutamine deamidase CheD (MCP methylation), and CheV (CheA docking to MCP) (14, 18).

FIG 1
FIG 1

Chemotaxis cluster organization in C. crescentus and growth using different carbon sources. (A) Schematic representation of the central chemotaxis apparatus. (B) Cell cycle-dependent transcriptional regulation of genes present in the major and alternate chemotaxis clusters. Data were extracted from previous global transcriptome analyses (27). The approximate C. crescentus cell cycle progression is shown at the top. RPKM, reads per kilobase million. (C) Genomic organization of the major and alternate chemotaxis clusters (gene locus numbers are given in Table 1). Genes encoding the central chemotaxis apparatus are represented using the same colors as in panel A; chemotaxis accessory proteins are represented in cyan. (D) Growth yield (OD600 after 24 h of incubation at 30°C) of CB15 WT grown using different carbon sources. The results are given as the mean from 3 independent replicates, and error bars represent the standard error of the mean (SEM).

In Caulobacter crescentus, the chemotaxis apparatus forms concomitantly with the flagellum apparatus (21, 22). This dimorphic bacterium starts its life as a piliated flagellated motile swarmer cell and then transitions to a sessile stalked cell by retracting its pili, shedding its flagellum, and synthesizing an adhesive holdfast followed by a stalk at the same pole (Fig. 1B). The chemoreceptor McpA is the best-characterized MCP in C. crescentus, and it is synthesized in a cell cycle-dependent manner, similarly to the other chemotaxis proteins encoded in this major chemotaxis cluster (21, 23–27) (Fig. 1B). McpA is synthesized at the new pole of predivisional cells; thus, newborn swarmer cells inherit McpA at the flagellar pole after division (28, 29). McpA is then degraded during the swarmer-to-stalked-cell transition (30) via proteolysis by ClpX (31), and this temporally regulated proteolysis plays an important role in the asymmetric distribution of McpA (30).

C. crescentus irreversibly adheres to surfaces and forms a biofilm by producing an adhesive holdfast (32–35). This polysaccharide adhesin contains β-1,4-N-acetylglucosamine residues (36, 37), and recent work suggests that its structure consists of a backbone of glucose, mannose, N-acetylglucosamine, and xylose residues, with branches at the C-6 position in the glucose and mannose residues (38). In addition to these sugar entities, holdfast contains peptides and DNA molecules (39). When cells are grown in complex medium, holdfast synthesis is temporally regulated via two pathways, a developmental program, or contact with a surface (1, 35). Newborn C. crescentus cells spend a portion of their life span as motile swarmer cells before differentiating though a highly controlled cell cycle progression into replicative stalked cells synthesizing holdfast and a stalk (Fig. 1B). However, swarmer cells that reach a surface bypass this developmental pathway and synthesize a holdfast within seconds of surface contact (40–43). The genes involved in holdfast synthesis and anchoring are transcribed in predivisional cells, resulting in newborn swarmer cells bearing complete and functional holdfast biosynthesis machinery. Holdfast production is regulated posttranslationally, and the second messenger molecule cyclic di-GMP (cdG) is the key regulator of holdfast production. Levels of intracellular cdG increase during the swarmer-to-stalked-cell differentiation, triggering the transition from motile to sessile states by flagellum shedding and holdfast production (44–46). In addition, HfsJ, a glycosyltransferase required for holdfast synthesis, directly binds cdG, triggering holdfast synthesis upon contact with the surface (43). The holdfast inhibitor HfiA also regulates holdfast synthesis by inhibiting HfsJ (47). The transcription of hfiA is regulated by cdG (48, 49) and the cell cycle progression, with transcript levels rising in predivisional cells and dropping in the swarmer-to-stalked cell transition (47). Environmental factors, such as blue light via the LovK-LovR system and nutrient availability, add to the control of hfiA expression (47, 50). Finally, HfiA is also regulated posttranscriptionally, as the chaperone DnaK affects HfiA levels, probably ensuring its stabilization in the cell (51). Overall, this multilayered control ensures that holdfast production and subsequent irreversible adhesion are tightly controlled at different levels.

In this study, we investigated the role of chemotaxis in biofilm formation and holdfast production in C. crescentus. While bacteria can swim toward gradients of nutrients, environmental stimuli, and signaling molecules via chemotaxis, they tend to follow their preferred growth substrates (16, 52). In this study, we decided to focus on molecules that can be metabolized by C. crescentus as carbon sources and that can also act as chemotaxis signals. As holdfast production and biofilm formation are regulated by nutrient availability in C. crescentus (47), our main aim was to determine if chemotaxis and cell adhesion were regulated via the same mechanism under specific nutrient growth conditions. We focused on dissecting the roles of both the major and alternate chemotaxis clusters in motility and adhesion. While previous works largely focus on the roles of genes present in the major chemotaxis cluster in swimming behaviors, here, we analyzed the role of the second alternate chemotaxis cluster by comparing mutants lacking the key CheA-type histidine kinases and the CheB-type methyltransferases that are encoded in the two chemotaxis clusters. We first show that only the major chemotaxis cluster is involved in chemotaxis, as only ΔcheAI and ΔcheBI mutants in the major cluster were unable to respond to a chemotactic gradient, while ΔcheAII and ΔcheBII mutants in the alternate cluster behaved like the wild type (WT). We then demonstrated that both clusters play a role in cell attachment and holdfast production in a complex nutrient-dependent manner. CheA and CheB proteins act antagonistically, and CheAI and CheAII positively regulate adhesion, while CheBI and CheBII repress it. These proteins also act to control the expression of the gene encoding the holdfast inhibitor HfiA. These results highlight different roles in regulating chemotaxis and biofilm formation for the two chemotaxis clusters.

RESULTS

Genomic organization of the chemotaxis genes in C. crescentus.There are two chemotaxis clusters encoded in the C. crescentus genome (53, 54). Historically, all mutations impairing the chemotactic response have been identified in a single cluster (28–30, 55), referred to as the major chemotaxis cluster (31). Hence, we named the second cluster the alternate chemotaxis cluster. The major cluster is cell cycle regulated (21, 22, 25–27, 56, 57), with a peak of expression occurring in predivisional cells (Fig. 1B). Transcripts from the alternate cluster are present at a significantly lower level than are those from the major locus. The genes encoding the MCPs in this cluster, mcpK and mcpG, are the only genes that seem to be cell cycle regulated, with a 1.5- to 2-fold increase in transcription in predivisional cells. All other genes present in the alternate locus are expressed at the same level throughout the cell cycle (Fig. 1B). This shows that the two loci are independently regulated and may fill different functions.

The two clusters are arranged similarly, with two MCPs, one CheA histidine kinase, three CheY response regulators, one CheW, one CheR, and one CheB (Fig. 1C). While the major cluster encodes a copy of accessory glutamine deamidase CheD (14) and the CheE protein of unknown function linked to chemotaxis (23, 25, 58), both are absent from the alternate cluster. In addition to the four MCP-encoding genes present in the two chemotaxis clusters, there are also 14 independent genes coding for putative MCPs (Table 1), suggesting that C. crescentus may sense a large array of specific attractants or repellents. The presence of a large number of MCPs in its genome is consistent with Caulobacter spp. being bacteria abundantly found in environments as diverse as oligotrophic freshwaters and nutrient-rich soils and being exposed to a wide array of attractants and repellent molecules (59, 60). No cheC, cheV, or cheZ homologs were detected in the C. crescentus genome (Table 1). There are several copies of key chemotaxis genes scattered in the genome, such as six copies of cheY, two of cheW, and one extra cheR (Table 1). Among the 12 homologs of cheY, five have been recently characterized as encoding CheY-like cdG effector (Cle) proteins (61). CleA is part of the major chemotaxis cluster, while CleB, CleC, CleD, and CleE are located independently in the genome (61) (Table 1). These Cle proteins bind to cdG and are involved in tuning of the flagellar motor activity, resulting in the subsequent increase of holdfast production upon contact with the surface (61).

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TABLE 1

Putative chemotaxis genes of C. crescentus

In this study, we investigated the behaviors of in-frame deletion mutants lacking either cheA (encoding the central histidine kinase that transduces the signal from the MCP receptor to the key response regulator protein CheY) or cheB (encoding the methyltransferase response regulator involved in removing a methyl group from the receptor MCP and modulating the chemotactic response). Genes present in the major chemotaxis cluster have been previously named cheAI and cheBI (62), so we named the genes present in the alternate chemotaxis cluster cheAII and cheBII. We constructed in-frame deletions of each of the cheA (cheAI and cheAII) and cheB (cheBI and cheBII) genes in C. crescentus CB15 as single- and double-deletion combinations.

Determination of carbon sources metabolized by C. crescentus.To determine the appropriate carbon sources to use in this study, we first surveyed a wide array of compounds that could be metabolized by C. crescentus as the sole carbon source. We tested a range of complex carbon, sugars, amino acids, organic acids, and alcohols. We monitored the growth yield obtained in M2 medium supplemented with a single given carbon source after 24 h and compared it to the growth yield and generation time in complex peptone-yeast extract (PYE) medium (Fig. 1D and Table 2). While M2-defined media provide inorganic phosphate, ammonium salts, and carbon, bacteria must de novo synthesize amino acids and nucleotides, which are crucial for growth. C. crescentus preferentially grew using complex carbon and sugars (Fig. 1D), confirming previous observations (59, 63). Mutations we made in the major and alternate chemotaxis clusters did not impair growth (Table 2; see also Fig. S1 in the supplemental material). To focus on carbon sources metabolized more efficiently by C. crescentus, we chose tryptone as an example of a complex carbon source and chose glucose, maltose, sucrose, and xylose as examples of sugars to conduct our studies. With the exception of sucrose, these sugars have been previously reported to be specific chemoattractant sugars for C. crescentus (55, 64).

View this table:
TABLE 2

Generation times of WT and mutant strains of C. crescentus CB15 grown using different carbon sources

Only the major chemotaxis cluster is involved in the chemotactic response toward different carbon sources.In C. crescentus, previous studies on chemotaxis almost exclusively focused on proteins encoded by the major chemotaxis cluster, with CheAI (CC_0433) (28, 64), CheBI (CC_0436) (28, 55), and CheRI (CC_0435) (28, 55) receiving most of the attention. A systematic study of all two-component signal transduction genes showed that CheAI and CheBI are important for swimming through semisolid plates containing complex PYE medium, while CheAII and CheBII are not (65). Finally, a more recent work investigated the chemotactic behavior of CheB and CheR null mutants (in-frame deletions of cheBI and cheBII or cheRI, cheRII, and cheRIII) toward galactose (66).

Here, we monitored the swimming behaviors of C. crescentus CB15 WT and mutant strains through semisolid agar plates containing different sugars as sole carbon sources (Fig. 2); only motile bacteria able to respond to a chemotactic gradient can form a ring under such conditions (55). Mutants in the alternate chemotaxis cluster (ΔcheAII and ΔcheBII) exhibited WT behavior, suggesting that the alternate cluster is not involved in chemotaxis (Fig. 2). However, both mutants in the major chemotaxis cluster (ΔcheAI and ΔcheBI) were impaired in chemotaxis, as deduced from their reduced ability to swim through semisolid medium compared to the WT (Fig. 2). Their defects could be complemented in trans by a replicating plasmid encoding a copy of cheAI or cheBI under the control of a constitutive promoter (Fig. S2). However, the cross-complementation with the paralogous gene could not restore the phenotype, as ΔcheAI and ΔcheBI mutants constitutively expressing in trans cheAII and cheBII, respectively, have swim rings similar to those of the mutants bearing the empty plasmid controls (Fig. S2).

FIG 2
FIG 2

Motility assays in semisolid agar. (A) Representative images of swim rings obtained after 5 days of incubation in semisolid plates made with M2 medium plus carbon source or PYE plus Noble agar (0.4%). Each image is 1 by 1 cm. (B) Swim diameters of the different strains using different carbon sources. Results are normalized to the WT ring diameter on the same plate type. Bar graphs indicate the mean from five independent replicates, and error bars represent the SEM. Statistical comparisons to the WT were calculated using paired t tests. ***, P < 0.001; **, P < 0.01; *, P < 0.05; ns, not significant.

The size of the swim ring, and therefore the amplitude of the chemotactic response, was different depending on the carbon source (Fig. 2); there was a 75% decrease for the ΔcheAI and ΔcheBI mutants when glucose, sucrose, or xylose was used as the carbon source, while a less drastic decrease was observed in the presence of other carbon sources (50% for maltose and 25% for tryptone and PYE media). The swimming behaviors of the double mutants confirm that the major chemotaxis cluster is the only one involved in chemotaxis under the conditions tested here. Both the ΔcheAI ΔcheAII and the ΔcheBI ΔcheBII mutants phenocopied the ΔcheAI and ΔcheBI single mutants and formed smaller swim rings than did the WT (Fig. 2). The ΔcheAI ΔcheBI mutant showed a decreased swim ring similar to those of the ΔcheAI and ΔcheBI single mutants, confirming that these two genes act in the same pathway to regulate chemotaxis (Fig. 2). Taken together, our results strongly suggest that only the major chemotaxis cluster is involved in chemotaxis in C. crescentus. It is possible, however, that the alternate chemotaxis cluster is used for chemotaxis under some conditions other than the ones tested in this study.

It is also worth noting that, under our experimental conditions, the mutants in the major chemotaxis cluster exhibited a moderately reduced swim ring compared to the WT (25% to 75% reduction depending the tested carbon source) (Fig. 2), while previous works reported more drastic reductions, similar to a nonmotile mutant (28, 55, 65). To verify these results, we tested other strains of CheA and CheB mutants described in the literature as chemotaxis deficient (65, 66); in our hands, all chemotaxis mutants were able to form larger swim rings than did a nonmotile ΔflgE mutant control, but still smaller than did the WT (Fig. S3A). The amplitude of the response was dependent on the carbon source and the agar used. Indeed, when plates were made using Bacto agar (214030; BD Difco), the chemotaxis mutants could form larger rings than when using Noble agar (0142-01; BD Difco). Interestingly, while M2 plates with no carbon source added did not support growth when made with Noble agar, some growth was noticeable using Bacto agar (Fig. S3B), suggesting that Bacto agar contains enough carbon traces to be metabolized by C. crescentus.

Both the major and alternate chemotaxis clusters are involved in biofilm regulation.As chemotaxis is involved in surface colonization and biofilm formation in different microorganisms, we tested our mutant strains for the ability bind to surfaces. We first quantified the amount of biofilm formed after 24 h under static conditions, using the five aforementioned carbon sources and PYE medium (Fig. S4). The ΔcheAI mutant was severely impaired in biofilm formation when grown in defined M2 medium, regardless of the tested carbon source. The amount of biofilm formed by the mutant corresponded to 30 to 50% of the WT levels, depending on the given carbon source (Fig. S4). Adhesion by the ΔcheAII mutant was reduced only when grown with certain carbon sources, as follows: while biofilm formation of the ΔcheAII mutant was not significantly different from that of the WT when grown with glucose, sucrose, or tryptone, it dropped to ΔcheAI mutant levels in M2 supplemented with maltose or xylose (Fig. S4). Intriguingly, both the ΔcheBI and ΔcheBII mutants formed more biofilm than did the WT when grown in defined medium (Fig. S4), suggesting that both CheB proteins are somehow involved in negatively regulating biofilm formation. Mutant phenotypes could be rescued by expressing a copy of the deleted gene on a replicating plasmid under the control of a constitutive promoter, but they could not be cross-complemented (Fig. S5).

To determine what stage of biofilm formation was impacted in these mutants, we monitored their attachment kinetics in the same static system. We focused on PYE and M2 supplemented with glucose (M2G) or xylose (M2X), since the adhesion phenotypes of the ΔcheAI and ΔcheAII mutants were different when grown in these different media. We first focused on cheA mutants (Fig. 3A). In PYE, the ΔcheAI mutant was impaired in biofilm initiation, with only 30% of the biomass attached compared to the WT strain after the first 10 h, but this strain eventually caught up and formed as much biofilm as did the WT at later time points, suggesting that this strain is impaired in early stages of biofilm formation only, when cells are grown in PYE (Fig. 3A, left). However, when glucose was used as the sole carbon source, the ΔcheAI mutant was affected in both biofilm initiation and maturation, as the amount of biomass attached was around 30% of the WT at each time point (Fig. 3A, middle). In both PYE and M2G media, ΔcheAII mutant adhesion was similar to that of the WT. Finally, in M2X, when xylose was the sole carbon source, both ΔcheAI and ΔcheAII mutants formed 50% less biofilm than did the WT (Fig. 3A, right). In all cases, the ΔcheAI ΔcheAII double mutant phenocopied the ΔcheAI single mutant (Fig. 3A). As CheAI is involved in chemotaxis (Fig. 2), we conclude that chemotaxis and biofilm formation pathways are interconnected through this crucial histidine kinase. In addition, even if CheAII is not involved in chemotaxis (Fig. 2), this protein plays a role in biofilm regulation. This regulation is carbon source specific and is not dependent on chemotaxis.

FIG 3
FIG 3

Biofilm formation over time. Amount of biofilm formed over time in PYE, M2 plus glucose (M2G), and M2 plus xylose (M2X) media. Cultures were grown in 24-well polystyrene plates, and the amount of biomass attached to the inside of the wells over time was quantified using crystal violet. Values are given as the average from crystal violet staining of triplicate samples of at least two independent experiments. The y axis error is represented as the SEM. (A) cheA mutants. (B) cheB mutants. (C) Major cluster mutants. (D) Minor cluster mutants.

We then assayed CheB mutants (Fig. 3B). In PYE, the amount of biofilm formed over time by the ΔcheBI and ΔcheBII mutants was similar to that of the WT (Fig. 3B, left), suggesting that these proteins are not involved in biofilm regulation in complex medium. However, in M2G and M2X, adhesion was more efficient in both CheB mutants than in the WT, with overall 1.5 to 2 times more cells attached to the surface at any given time point (Fig. 3B). The amount of biofilm formed by the ΔcheBI ΔcheBII double mutant was similar to that of each single mutant. Interestingly, the ΔcheAI ΔcheBI and ΔcheAII ΔcheBII double mutants both exhibited a hyperbiofilm phenotype, with an increased adhesion similar to the cheB single mutants (Fig. 3C and D). This suggests that CheB acts downstream of CheA in regulating adhesion in C. crescentus.

Taken together, these results show an intriguing relationship between chemotaxis and biofilm regulation. CheA and CheB act in opposition; while mutations in cheA cause a decrease in adhesion, cheB deletions cause an increase in adhesion. Although both CheAI and CheBI have been shown to be crucial for chemotaxis, these proteins have distinct antagonistic roles in biofilm regulation. In addition, these data highlight a role for the alternate chemotaxis cluster in the regulation of biofilm formation through CheAII and CheBII. Like their major chemotaxis cluster counterparts, CheAII and CheBII act in an opposite manner but do so conditionally, since CheAII is involved in biofilm regulation only in the presence of certain carbon sources.

CheA and CheB proteins regulate holdfast production in an antagonistic manner.The adhesive holdfast is essential for long-term adhesion in C. crescentus (32–34). Because long-term adhesion was altered in the tested Che mutants, we sought to determine whether changes in holdfast production could be responsible for this defect. To quantify the proportion of single cells harboring a holdfast in the mixed population, we stained early exponential cultures grown in PYE, M2G, or M2X with fluorescent wheat germ agglutinin (WGA), which specifically stains the N-acetylglucosamine residues present in the holdfast (37), and quantified the number of cells with a holdfast by fluorescence microscopy.

As previously reported, the number of cells harboring a holdfast is drastically different when the cells are grown in complex PYE medium than when grown in nutrient-defined M2X (47) or M2G (48) medium. Interestingly, there was also a difference in holdfast formation depending on the carbon source used in the growth medium, as around 20% of WT cells harbored a holdfast when grown in the presence of glucose, while less than 10% did in the presence of xylose (Fig. 4A). In all tested media, the number of cells producing a holdfast in the ΔcheAI mutant population was reduced by half compared to the WT (Fig. 4A). In PYE, the number of holdfasts detected in the other tested mutants did not significantly differ from the WT population (Fig. 4A). However, in defined media, both of the ΔcheB mutant populations produced approximately 20% more holdfasts than did the WT (Fig. 4A). The number of holdfasts present in the ΔcheAII mutant phenocopied the WT in M2G and the ΔcheAI mutant strain in M2X (Fig. 4A). These results are in agreement with the biofilm phenotypes presented in Fig. 3 and S4.

FIG 4
FIG 4

Role of the holdfast inhibitor HfiA. (A) Quantification of cells harboring a holdfast in mixed populations. Cells were stained using Alexa Fluor 488-WGA and imaged by fluorescence microscopy. The results represent the average from three independent replicates (more than 300 cells per replicate), and the error bars represent the SEM. (B) β-Galactosidase activity of PhfiA-lacZ transcriptional fusions in PYE, M2G, and M2X media supplemented with tetracycline. The results represent the average from 9 independent cultures (assayed on 3 different days), and the error bars represent the SEM. Statistical comparisons to the WT were calculated using unpaired t tests. (C) Biofilm formation after 24 h of incubation at 30°C in PYE and M2 media supplemented with glucose or xylose. The results are normalized to the WT biofilm formation in the given medium. Error bars represent the SEM from three independent replicates run in duplicate. Statistical comparisons to the WT were calculated using unpaired t tests. ***, P < 0.001; **, P < 0.01; *, P < 0.05; ns, not significant.

Transcription of the holdfast inhibitor-encoding gene, hfiA, is regulated by CheA and CheB.In C. crescentus, holdfast production is regulated by nutrient availability via the holdfast inhibitor protein HfiA (47). In defined M2G and M2X media, hfiA expression is upregulated, resulting in a significant decrease in holdfast production compared to cells grown in PYE (47, 48). To determine if HfiA was playing a role in the differences in holdfast production observed in the present study, we measured the expression of hfiA using lacZ transcriptional fusions in WT and the chemotaxis mutants. In PYE, the activity of the hfiA promoter was minimal. Still, hfiA expression in the ΔcheAI mutant was approximately 50% higher than that in the other tested strains (Fig. 4B). In defined M2 media, we observed a drastic increase in hfiA transcription compared to in PYE, as previously reported (47–49) (Fig. 4B). There was also an overall increase in hfiA expression in M2X compared to that in M2G. In defined M2 media, hfiA expression was elevated in the ΔcheAI strain while it was decreased in both ΔcheB mutants (Fig. 4B). These results correlate with holdfast quantification (Fig. 4A); hfiA expression was lower in populations that have more cells with a holdfast and tend to form more biofilms.

We also looked at how an hfiA deletion or overexpression would affect the tested Che mutants. We first quantified biofilm formation in hfiA che mutants (Fig. 4C). In PYE medium, the ΔhfiA mutant produced slightly more biofilm than did the WT, as expected. All cheA and cheB single mutants produced similar amounts of biofilm compared to the WT strain, and the hfiA che double mutants all phenocopied the ΔhfiA mutant in this complex medium. In M2G and M2X defined media, the double mutants also phenocopied the ΔhfiA mutant. Based on these results, we conclude that the che genes and hfiA are acting in the same pathway to regulate holdfast production. Biofilm formation by strains where hfiA is chromosomally inserted at the xylX locus in the che mutants and induced shows a reduction of 80 to 90% compared to the WT empty vector strain (Fig. S6). This result further suggests that hfiA acts downstream of the holdfast regulation driven by the che proteins. Overall, these observations show that chemotaxis genes regulate hfiA expression and thereby control holdfast production in response to the carbon sources available in the medium.

DISCUSSION

In this work, we investigated the roles of the two C. crescentus chemotaxis clusters in chemotaxis and surface attachment. We showed that only the major cluster is involved in chemotaxis, while both clusters regulate biofilm formation and holdfast production. Our results support a model where both CheAI and CheAII proteins negatively regulate the expression of the gene encoding the holdfast inhibitor protein, HfiA, while CheB proteins activate its expression in response to the carbon source present in the medium. It has been recently shown that disturbances in flagellum or pilus synthesis modify holdfast production via hfiA regulation (48, 49), and we now add chemotaxis proteins as regulators of holdfast synthesis via HfiA.

Other chemotaxis proteins, specifically, the CheY-like Cle proteins, have been shown to be involved in chemotaxis and holdfast synthesis (61). Interestingly, only CleA, located in the major chemotaxis cluster, has been shown to play a role in chemotaxis regulation, confirming our observations that only the major chemotaxis cluster properly functions as a regulator of chemotaxis (61). In addition, Cle proteins are involved in the regulation of holdfast production upon surface contact (61). However, holdfast synthesis by surface contact stimulation does not occur in defined M2 medium (48), suggesting that the regulation observed in our work is linked to developmentally programmed holdfast production and therefore occurs via a different mechanism. The multifunctional response regulator MrrA, which is essential for the general stress response in C. crescentus, has been recently shown to play a role in chemotaxis and holdfast production via the modulation of hfiA expression (67). It could be interesting to test if this global regulator is involved in holdfast regulation by the Che proteins. Another putative player to explore in the future is cdG. Indeed, this ubiquitous messenger molecule is involved in chemotaxis (61), holdfast production (44, 45), and hfiA expression (48, 49), and it could be involved in the modulation of holdfast synthesis by the chemotaxis proteins.

Many studies in other bacteria have shown that chemotaxis-like clusters are involved in behaviors independent from chemotaxis, such as cell differentiation or biofilm formation (3, 68). For example, regulatory pathways homologous to the chemotaxis system have been shown to control cyst formation in Rhodospirillum centenum (69), fibril polysaccharide production in Myxococcus xanthus (70), and cell aggregation and biofilm formation in Azospirillum brasilense (11). In these species, different proteins within the chemotaxis-like systems act antagonistically. In R. centenum, MCP, CheW, CheR, and CheA positively regulate cyst formation, while CheY and CheB are negative regulators (69). In M. xanthus, the chemotaxis-like Dif pathway regulates the formation of fibril polysaccharide, a crucial component for fruiting body and spore formation. Within the Dif operon, DifD (CheY homolog) and DifG (CheC phosphatase homolog) are negative regulators of fibril production, whereas DifA (MCP homolog), DifC (CheW homolog), and DifE (CheA homolog) are positive regulators (70–72). In A. brasilense, CheA and CheY repress exopolysaccharide (EPS) production involved in cell-cell aggregation and biofilm formation, while CheB and CheR enhance it (11). Our results show that in C. crescentus, different Che proteins also have antagonistic effects on holdfast polysaccharide production, with CheA and CheB proteins from both clusters acting as positive and negative regulators, respectively.

The best-characterized chemotaxis-like operon involved in biofilm formation is the Wsp system of Pseudomonas aeruginosa. This system controls EPS production by modulating cdG levels in response to contact with a surface (73, 74). Briefly, WspR is a CheY homolog and a diguanylate cyclase that acts as the final response regulator of the Wsp system (73, 75). In its active form, WspR produces cdG, which in turn activates the production of Pel and Psl exopolysaccharides and biofilm formation. In that system, WspF (CheB homolog) is a modulator of WpsR. The deletion of wpsF results in increased phosphorylation of WspR, which negatively regulates the polysaccharides Psl and Pel while interfering with the intracellular levels of cdG (73, 75). Future work will determine if holdfast regulation by CheA/CheB in C. crescentus involves a similar mechanism.

In conclusion, we have demonstrated that the two chemotaxis clusters of C. crescentus have distinct roles. Our data show that while the major cluster is involved in both chemotaxis and holdfast production, the alternate cluster is a chemotaxis-like system involved in holdfast regulation but not chemotaxis toward the compounds tested. Mutants lacking the kinases CheAI and CheAII are impaired in cell attachment, resulting from a defect in holdfast production, while CheBI and CheBII mutants produce more holdfasts and form more robust biofilms. We also showed that the regulation of holdfast synthesis by Che proteins is due to the modulation of the expression of hfiA, encoding the holdfast inhibitor protein HfiA. These data suggest a model where CheA proteins promote holdfast synthesis, while CheB proteins repress it, by modulating hfiA expression. Further identification of players in this regulatory pathway and more in-depth exploration of the mechanism by which this occurs may reveal how bacteria respond to external stimuli to optimize bacterial adhesion and surface colonization in various environments.

MATERIALS AND METHODS

Bacterial strains, plasmids, and growth conditions.The bacterial strains used in this study are listed in Table S1 in the supplemental material. C. crescentus strains were grown at 30°C in defined M2 medium (76) supplemented with 0.2% (wt/vol) of a given carbon source (listed in Table 2) or in complex peptone-yeast extract (PYE) medium (59). When appropriate, 5 μg/ml kanamycin or 1 μg/ml tetracycline was added to the medium. For induction of the vanillate (Van) promoter in pMT630-derivative constructs, 0.5 mM vanillate was added to the cultures before inoculation and incubation. Escherichia coli Silver α select cells (Bioline) were used for cloning and were grown in LB medium at 37°C with 25 μg/ml kanamycin or 10 μg/ml tetracycline when appropriate.

In-frame deletion mutants were obtained by double-homologous recombination, as previously described (77). Briefly, PCRs were performed using C. crescentus CB15 genomic DNA (gDNA) as the template to amplify 500-bp fragments from the upstream and downstream regions of the gene to be deleted. The primers designed for these in-frame deletions are listed in Table S3. PCR fragments were gel purified using the Zymoclean gel DNA recovery kit (Zymo Research) and then digested by BamHI and XhoI or XhoI and HindIII for upstream or downstream fragments, respectively. Purified digested fragments were then cloned into the suicide vector pNPTS138 that had been digested by BamHI and HindIII. The pNPTS138-based constructs were transformed into E. coli Silver α select cells and then introduced into C. crescentus by electroporation. The two-step recombination was carried out first by selecting integrants on PYE supplemented with kanamycin and second by growing them overnight without selection (into 5 ml liquid PYE at 26˚C) and plating the overnight cultures (1 μl) on PYE supplemented with 3% sucrose to select for bacteria that lost the plasmid as part of a second recombination event (77). Then, the mutants were checked by sequencing to confirm the presence of the deletion.

The complementation plasmids, harboring cheAI, cheAII, cheBI, or cheBII, were constructed as follows. C. crescentus CB15 gDNA was used as the template to PCR amplify the genes of interest using primers containing HindIII (forward primers) and KpnI (downstream primers) restriction sites (Table S2). PCR products were gel purified using the Zymoclean gel DNA recovery kit (Zymo Research), digested using HindIII and KpnI, and ligated into plasmid pMR10 (78), extracted using the Zippy Plasmid prep kit (Zymo Research), and digested by the same enzymes.

Growth curves and generation time calculations.Bacterial growth in the different media was measured in 3-ml liquid cultures (in 15-ml glass tubes) with shaking at 300 rpm. Overnight cultures were diluted in the same culture medium to an optical density at 600 nm (OD600) of 0.05 and incubated for 24 h. The OD600 was measured at various time intervals to generate growth curves (OD600 versus time). Generation times were calculated from the exponential part of the growth curves using the single exponential-growth function in the GraphPad Prism 6 software.

Motility assays in semisolid media.Motility assays were performed using semisolid agar plates. Plates were poured using PYE or M2 medium supplemented with 0.2% (wt/vol) of the appropriate carbon source and 0.4% (wt/vol) Noble agar (reference 0142-01; Difco). Cells were stabbed in the soft agar and incubated in a humid chamber at 30°C for 5 days. The diameter of the swimming ring formed by each tested strain was measured manually.

Biofilm assays.Biofilm assays in multiwell plates were performed using two different setups that yield similar results (79), as follows: (i) adhesion to polyvinyl chloride (PVC) microscope coverslips placed vertically in plastic 12-well plates or (ii) adhesion to the inside surface of the wells of untreated plastic 24-well plates. Bacteria were grown to mid-log phase (OD600, 0.4 to 0.8) in the chosen medium and diluted to an OD600 of 0.05 in the same medium in 3 or 0.5 ml for the 12- or 24-well plate setup, respectively. Plates were incubated at 30°C for different times. Biofilms attached to coverslips or inside surfaces of the wells were quantified, as follows: wells or coverslips were rinsed with distilled H2O (dH2O) to remove nonattached bacteria, stained using 0.1% crystal violet (CV), and rinsed again with dH2O. The CV from the stained attached biomass was eluted using 10% (vol/vol) acetic acid and was quantified by measuring the absorbance at 600 nm (A600). Biofilm formation was normalized to A600/OD600 and expressed as a ratio of the WT level.

Holdfast quantification using fluorescently labeled WGA lectin.The number of cells harboring a holdfast in mixed populations was quantified by fluorescence microscopy. Holdfasts were detected with Alexa Fluor 488-WGA. Early exponential-phase cultures (OD600, 0.2 to 0.4) were mixed with Alexa Fluor 488-WGA (0.5 μg/ml final concentration). One microliter of WGA-stained cells was spotted on a 1.5-mm glass coverslip and covered with an agarose pad (1% SeaKem LE agarose dissolved in dH2O). Holdfasts were imaged by epifluorescence microscopy using a Nikon Ti-2 microscope with a Plan Apo 60× objective, a green fluorescent protein (GFP)/DsRed filter cube, a Hamamatsu Orca Flash 4.0 camera, and the Nikon NIS Elements imaging software. The number of individual cells with a holdfast was calculated manually from microscopy images in the Nikon NIS Elements imaging software.

β-Galactosidase assays.Strains bearing the transcriptional reporter plasmid of the hfiA gene promoter fused to lacZ (47) were inoculated from freshly grown colonies into 5 ml of a chosen medium containing 1 μg/ml tetracycline and were then incubated at 30°C overnight. Cultures were then diluted in the same culture medium to an OD600 of 0.05 and incubated until an OD600 of 0.15 to 0.25 was reached. β-Galactosidase activity was measured colorimetrically, as described previously (80). A volume of 200 μl of culture was mixed with 600 μl of Z buffer (60 mM Na2HPO4, 40 mM NaH2PO4, 10 mM KCl, 1 mM MgSO4, 50 mM β-mercaptoethanol), 50 μl of chloroform, and 25 μl of 0.1% SDS. Two hundred microliters of the substrate o-nitrophenyl-β-d-galactopyranoside (4 mg/ml) was then added to the cell mixture, and the time until development of a yellow color was recorded. The reaction was stopped by adding 400 μl of 1 M Na2CO3 to raise the pH to 11. A420 was measured, and the Miller units of β-galactosidase activity were calculated as (A420 × 1,000)/[(OD600 × t) × v], where t is the incubation time in minutes and v is the volume of culture (in milliliters) used in the assay. The β-galactosidase activity of WT CB15/plac290 (empty vector control) was used as a blank sample reference.

ACKNOWLEDGMENTS

We thank Aretha Fiebig, Mike Laub, and Martin Thanbichler for providing strains, as well as the members of the Brun laboratory and S. Zappa for critical reading of the manuscript.

This study was supported by grant R35GM122556 from the National Institutes of Health and by a Canada 150 Research Chair in Bacterial Cell Biology to Y.V.B.

FOOTNOTES

    • Received 22 January 2019.
    • Accepted 16 May 2019.
    • Accepted manuscript posted online 20 May 2019.

  • Supplemental material for this article may be found at https://doi.org/10.1128/JB.00071-19.

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KEYWORDS

Caulobacter crescentus
bacterial adhesion
biofilms
chemotaxis
holdfast

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