The Two Chemotaxis Clusters in Caulobacter crescentus PlayDifferent Roles in Chemotaxis and Biofilm Regulation

Cécile Berne,a,b Yves V. Bruna,b

aDepartment of Biology, Indiana University, Bloomington, Indiana, USAbDépartement de Microbiologie, Infectiologie et Immunologie, Université de Montréal, Montréal, Québec, Canada

ABSTRACT The holdfast polysaccharide adhesin is crucial for irreversible cell adhe-sion and biofilm formation in Caulobacter crescentus. Holdfast production is tightlycontrolled via developmental regulators, as well as via environmental and physicalsignals. Here, we identify a novel mode of regulation of holdfast synthesis that involveschemotaxis proteins. We characterized the two identified chemotaxis clusters of C.crescentus and showed that only the previously characterized major cluster is in-volved in the chemotactic response toward different carbon sources. However, bothchemotaxis clusters encoded in the C. crescentus genome play a role in biofilm for-mation and holdfast production by regulating the expression of hfiA, the gene en-coding the holdfast inhibitor HfiA. We show that CheA and CheB proteins act in anantagonistic manner, as follows: while the two CheA proteins negatively regulatehfiA expression, the CheB proteins are positive regulators, thus providing a modula-tion of holdfast synthesis and surface attachment.

IMPORTANCE Chemosensory systems constitute major signal transduction pathwaysin bacteria. These systems are involved in chemotaxis and other cell responses toenvironment conditions, such as the production of adhesins to enable irreversibleadhesion to a surface and surface colonization. The C. crescentus genome encodes twocomplete chemotaxis clusters. Here, we characterized the second novel chemotaxis-likecluster. While only the major chemotaxis cluster is involved in chemotaxis, both che-motaxis systems modulate C. crescentus adhesion by controlling expression of theholdfast 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 for-mation of biofilms in response to the environment.

KEYWORDS Caulobacter crescentus, bacterial adhesion, biofilms, chemotaxis, holdfast

In their natural habitat, most bacteria are organized in complex surface-associatedmulticellular 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 withthe surface. The cells then divide to form multicellular microcolonies, which even-tually develop into a mature biofilm (1). Communal life on a surface is believed tobe beneficial, as it provides protection from predators and xenobiotic stresses (2). Theenvironment at the surface is highly heterogenous, with the presence of variouscompounds adsorbed on the surface and the formation of gradients near it (1). Toinitiate attachment, bacteria must approach the surface either by passive transport orby active swimming (1). Both active swimming toward the surface and initial surfaceattachment 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 andin cell-cell aggregation (11, 12). Finally, chemotaxis is also involved in later stages ofbiofilm formation, as is the case for single Pseudomonas aeruginosa cells that can

Citation Berne C, Brun YV. 2019. The twochemotaxis clusters in Caulobacter crescentusplay different roles in chemotaxis and biofilmregulation. J Bacteriol 201:e00071-19. https://doi.org/10.1128/JB.00071-19.

Editor George O’Toole, Geisel School ofMedicine at Dartmouth

Copyright © 2019 American Society forMicrobiology. All Rights Reserved.

Address correspondence to Yves V. Brun,yves.brun@umontreal.ca.

Received 22 January 2019Accepted 16 May 2019

Accepted manuscript posted online 20 May2019Published

MEETING PRESENTATION

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actively respond to a chemical gradient and subsequently reposition themselves on thesurface within a mature biofilm (13).

The central chemotaxis system is composed of a chemoreceptor, or methyl-acceptingchemotaxis protein (MCP), and CheW, CheA, and CheY proteins (14–18). These proteinsform large complexes of hexagonally packed arrays, localized at the cell pole, withMCPs being transmembrane proteins, and with CheW, CheA, and CheY being locatedin the cytoplasm (19, 20). The chemotactic signal is sensed by the MCP and transducedto the sensory histidine kinase CheA via the scaffolding protein CheW (Fig. 1A). If the

FIG 1 Chemotaxis cluster organization in C. crescentus and growth using different carbon sources. (A) Schematic representation of the central chemotaxisapparatus. (B) Cell cycle-dependent transcriptional regulation of genes present in the major and alternate chemotaxis clusters. Data were extracted fromprevious 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 chemotaxisapparatus are represented using the same colors as in panel A; chemotaxis accessory proteins are represented in cyan. (D) Growth yield (OD600 after 24 h ofincubation 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 representthe standard error of the mean (SEM).

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signal is an attractant, CheA autophosphorylation is inhibited, while a repellant signalactivates it. Phosphorylated CheA (CheA�P) serves as a phosphodonor to the CheY andCheB response regulators. CheY�P interacts with the flagellum apparatus, modulatingflagellum rotation and swimming behavior. CheB�P removes a methyl group from theMCP, reducing the overall activity of the signal transduction cascade and modulatingthe chemotactic response. Optional accessory proteins have also been linked to thechemotaxis apparatus and play auxiliary roles in chemotaxis regulation, such as thephosphatases CheC, its homolog CheX, and CheZ (CheY�P hydrolyzation), the glu-tamine deamidase CheD (MCP methylation), and CheV (CheA docking to MCP) (14, 18).

In Caulobacter crescentus, the chemotaxis apparatus forms concomitantly with theflagellum apparatus (21, 22). This dimorphic bacterium starts its life as a piliated flagellatedmotile 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 atthe 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 otherchemotaxis 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 cellsinherit McpA at the flagellar pole after division (28, 29). McpA is then degraded duringthe swarmer-to-stalked-cell transition (30) via proteolysis by ClpX (31), and this tem-porally regulated proteolysis plays an important role in the asymmetric distribution ofMcpA (30).

C. crescentus irreversibly adheres to surfaces and forms a biofilm by producingan adhesive holdfast (32–35). This polysaccharide adhesin contains �-1,4-N-acetyl-glucosamine residues (36, 37), and recent work suggests that its structure consists of abackbone of glucose, mannose, N-acetylglucosamine, and xylose residues, withbranches at the C-6 position in the glucose and mannose residues (38). In addition tothese sugar entities, holdfast contains peptides and DNA molecules (39). When cells aregrown in complex medium, holdfast synthesis is temporally regulated via two path-ways, a developmental program, or contact with a surface (1, 35). Newborn C. crescentuscells spend a portion of their life span as motile swarmer cells before differentiatingthough a highly controlled cell cycle progression into replicative stalked cells synthe-sizing holdfast and a stalk (Fig. 1B). However, swarmer cells that reach a surface bypassthis developmental pathway and synthesize a holdfast within seconds of surfacecontact (40–43). The genes involved in holdfast synthesis and anchoring are transcribedin predivisional cells, resulting in newborn swarmer cells bearing complete and func-tional holdfast biosynthesis machinery. Holdfast production is regulated posttransla-tionally, and the second messenger molecule cyclic di-GMP (cdG) is the key regulatorof holdfast production. Levels of intracellular cdG increase during the swarmer-to-stalked-cell differentiation, triggering the transition from motile to sessile states byflagellum shedding and holdfast production (44–46). In addition, HfsJ, a glycosyltrans-ferase required for holdfast synthesis, directly binds cdG, triggering holdfast synthesisupon contact with the surface (43). The holdfast inhibitor HfiA also regulates holdfastsynthesis 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 anddropping in the swarmer-to-stalked cell transition (47). Environmental factors, such asblue light via the LovK-LovR system and nutrient availability, add to the control of hfiAexpression (47, 50). Finally, HfiA is also regulated posttranscriptionally, as the chaperoneDnaK affects HfiA levels, probably ensuring its stabilization in the cell (51). Overall, thismultilayered control ensures that holdfast production and subsequent irreversibleadhesion are tightly controlled at different levels.

In this study, we investigated the role of chemotaxis in biofilm formation andholdfast production in C. crescentus. While bacteria can swim toward gradients ofnutrients, environmental stimuli, and signaling molecules via chemotaxis, they tend tofollow their preferred growth substrates (16, 52). In this study, we decided to focus onmolecules that can be metabolized by C. crescentus as carbon sources and that can alsoact as chemotaxis signals. As holdfast production and biofilm formation are regulated

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by nutrient availability in C. crescentus (47), our main aim was to determine if che-motaxis and cell adhesion were regulated via the same mechanism under specificnutrient growth conditions. We focused on dissecting the roles of both the major andalternate chemotaxis clusters in motility and adhesion. While previous works largelyfocus on the roles of genes present in the major chemotaxis cluster in swimmingbehaviors, here, we analyzed the role of the second alternate chemotaxis cluster bycomparing mutants lacking the key CheA-type histidine kinases and the CheB-typemethyltransferases that are encoded in the two chemotaxis clusters. We first show thatonly the major chemotaxis cluster is involved in chemotaxis, as only �cheAI and �cheBImutants in the major cluster were unable to respond to a chemotactic gradient, while�cheAII and �cheBII mutants in the alternate cluster behaved like the wild type (WT).We then demonstrated that both clusters play a role in cell attachment and holdfastproduction in a complex nutrient-dependent manner. CheA and CheB proteins actantagonistically, and CheAI and CheAII positively regulate adhesion, while CheBI andCheBII repress it. These proteins also act to control the expression of the gene encodingthe holdfast inhibitor HfiA. These results highlight different roles in regulating che-motaxis and biofilm formation for the two chemotaxis clusters.

RESULTSGenomic organization of the chemotaxis genes in C. crescentus. There are two

chemotaxis clusters encoded in the C. crescentus genome (53, 54). Historically, allmutations 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 thesecond 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 arethose from the major locus. The genes encoding the MCPs in this cluster, mcpK andmcpG, are the only genes that seem to be cell cycle regulated, with a 1.5- to 2-foldincrease in transcription in predivisional cells. All other genes present in the alternatelocus are expressed at the same level throughout the cell cycle (Fig. 1B). This shows thatthe 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 themajor cluster encodes a copy of accessory glutamine deamidase CheD (14) and theCheE protein of unknown function linked to chemotaxis (23, 25, 58), both are absentfrom the alternate cluster. In addition to the four MCP-encoding genes present in thetwo 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 orrepellents. The presence of a large number of MCPs in its genome is consistent withCaulobacter spp. being bacteria abundantly found in environments as diverse asoligotrophic freshwaters and nutrient-rich soils and being exposed to a wide array ofattractants and repellent molecules (59, 60). No cheC, cheV, or cheZ homologs weredetected in the C. crescentus genome (Table 1). There are several copies of keychemotaxis genes scattered in the genome, such as six copies of cheY, two of cheW, andone extra cheR (Table 1). Among the 12 homologs of cheY, five have been recentlycharacterized as encoding CheY-like cdG effector (Cle) proteins (61). CleA is part of themajor chemotaxis cluster, while CleB, CleC, CleD, and CleE are located independently inthe genome (61) (Table 1). These Cle proteins bind to cdG and are involved in tuningof the flagellar motor activity, resulting in the subsequent increase of holdfast produc-tion upon contact with the surface (61).

In this study, we investigated the behaviors of in-frame deletion mutants lackingeither cheA (encoding the central histidine kinase that transduces the signal fromthe MCP receptor to the key response regulator protein CheY) or cheB (encoding themethyltransferase response regulator involved in removing a methyl group from thereceptor MCP and modulating the chemotactic response). Genes present in the majorchemotaxis cluster have been previously named cheAI and cheBI (62), so we named the

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genes present in the alternate chemotaxis cluster cheAII and cheBII. We constructedin-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 determinethe appropriate carbon sources to use in this study, we first surveyed a wide array ofcompounds that could be metabolized by C. crescentus as the sole carbon source. Wetested a range of complex carbon, sugars, amino acids, organic acids, and alcohols. Wemonitored the growth yield obtained in M2 medium supplemented with a single givencarbon source after 24 h and compared it to the growth yield and generation time incomplex peptone-yeast extract (PYE) medium (Fig. 1D and Table 2). While M2-definedmedia provide inorganic phosphate, ammonium salts, and carbon, bacteria must denovo synthesize amino acids and nucleotides, which are crucial for growth. C. crescentuspreferentially grew using complex carbon and sugars (Fig. 1D), confirming previousobservations (59, 63). Mutations we made in the major and alternate chemotaxis

TABLE 1 Putative chemotaxis genes of C. crescentus

Cluster and gene Locus tag in CB15 Locus tag in NA1000 Gene product

Major chemotaxis clustermcpB CC_0428 CCNA_00437 Receptor MCPmcpA CC_0430 CCNA_00439 Receptor MCPcheY CC_0432 CCNA_00441 Response regulator protein CheYcheA CC_0433 CCNA_00442 Chemotaxis sensory histidine kinase CheAcheW CC_0434 CCNA_00443 Receptor binding protein CheWcheR CC_0435 CCNA_00444 Chemotaxis protein methyltransferase CheRcheB CC_0436 CCNA_00445 Chemotaxis protein methyltransferase CheBcheY CC_0437 CCNA_00446 Response regulator protein CheYcheD CC_0438 CCNA_00447 Chemotaxis protein CheDcleA CC_0440 CCNA_00449 CheY-like cdG effector CleA

Alternate chemotaxis clustercheY CC_0588 CCNA_00625 Response regulator protein CheYmcpG CC_0589 CCNA_00626 Receptor MCPcheY CC_0591 CCNA_00628 Response regulator protein CheYmcpK CC_0593 CCNA_00629 Receptor MCPcheA CC_0594 CCNA_00630 Chemotaxis sensory histidine kinase CheAcheW CC_0595 CCNA_00631 Receptor binding protein CheWcheY CC_0596 CCNA_00632 Response regulator protein CheYcheB CC_0597 CCNA_00633 Chemotaxis protein methyltransferase CheBcheR CC_0598 CCNA_00634 Chemotaxis protein methyltransferase CheR

Single genesmcpQ CC_0066 CCNA_00064 Receptor MCPmcpC CC_0343 CCNA_00160 Receptor MCPmcpN CC_0504 CCNA_00538 Receptor MCPcheW CC_0764 CCNA_00803 Receptor binding protein CheWcleB CC_1364 CCNA_01426 CheY-like cdG effector CleBmcpP CC_1399 CCNA_01465 Receptor MCPmcpD CC_1655 CCNA_01727 Receptor MCPcleC CC_2249 CCNA_02332 CheY-like cdG effector CleCmcpE CC_2281 CCNA_02364 Receptor MCPmcpM CC_2317 CCNA_02402 Receptor MCPmcpF CC_2691 CCNA_02773 Receptor MCPmcp CC_2810 CCNA_02901 Receptor MCPmcpO CC_2842 CCNA_02935 Receptor MCPmcpI CC_2847 CCNA_02940 Receptor MCPcheW CC_3025 CCNA_03120 Receptor binding protein CheWcleD CC_3100 CCNA_03198 CheY-like cdG effector CleDmcpJ CC_3145 CCNA_03247 Receptor MCPcleE CC_3155 CCNA_03257 CheY-like cdG effector CleEcheY CC_3258 CCNA_03367 Response regulator protein CheYmcpH CC_3349 CCNA_03459 Receptor MCPmcpL CC_3358 CCNA_03468 Receptor MCPcheY CC_3471 CCNA_03585 Response regulator protein CheYcheR CC_3472 CCNA_03586 Chemotaxis protein methyltransferase CheR

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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 chosetryptone 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 exceptionof sucrose, these sugars have been previously reported to be specific chemoattractantsugars for C. crescentus (55, 64).

Only the major chemotaxis cluster is involved in the chemotactic responsetoward different carbon sources. In C. crescentus, previous studies on chemotaxisalmost exclusively focused on proteins encoded by the major chemotaxis cluster, withCheAI (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 signaltransduction genes showed that CheAI and CheBI are important for swimming throughsemisolid plates containing complex PYE medium, while CheAII and CheBII are not (65).Finally, a more recent work investigated the chemotactic behavior of CheB and CheRnull mutants (in-frame deletions of cheBI and cheBII or cheRI, cheRII, and cheRIII) towardgalactose (66).

Here, we monitored the swimming behaviors of C. crescentus CB15 WT and mutantstrains 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 ringunder such conditions (55). Mutants in the alternate chemotaxis cluster (ΔcheAII andΔcheBII) exhibited WT behavior, suggesting that the alternate cluster is not involved inchemotaxis (Fig. 2). However, both mutants in the major chemotaxis cluster (ΔcheAI andΔcheBI) were impaired in chemotaxis, as deduced from their reduced ability to swimthrough semisolid medium compared to the WT (Fig. 2). Their defects could becomplemented in trans by a replicating plasmid encoding a copy of cheAI or cheBIunder the control of a constitutive promoter (Fig. S2). However, the cross-complementationwith the paralogous gene could not restore the phenotype, as �cheAI and �cheBI mutantsconstitutively expressing in trans cheAII and cheBII, respectively, have swim rings similarto those of the mutants bearing the empty plasmid controls (Fig. S2).

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

TABLE 2 Generation times of WT and mutant strains of C. crescentus CB15 grown using different carbon sources

Carbon source

Generation time (min) for:

WT �cheAI mutant �cheBI mutant �cheAII mutant �cheBII mutant

Complex carbon sourcesPYE 94 � 5 95 � 5 97 � 5 93 � 5 95 � 8Casamino Acids 123 � 5 122 � 7 116 � 8 125 � 5 119 � 7Peptone 115 � 5 128 � 4 110 � 5 118 � 5 125 � 8Tryptone 109 � 3 102 � 5 115 � 6 113 � 6 105 � 7

SugarsGlucose 121 � 6 125 � 8 122 � 8 118 � 5 120 � 8Maltose 125 � 9 127 � 7 132 � 8 129 � 7 131 � 9Sucrose 137 � 3 141 � 6 144 � 6 140 � 1 138 � 5Xylose 138 � 7 138 � 6 138 � 5 134 � 6 135 � 6

Amino acidsAlanine 473 � 10 496 � 10 487 � 9 500 � 10 478 � 9Glutamic acid 443 � 8 437 � 9 438 � 7 449 � 9 444 � 10

Organic acidsCitrate 573 � 17 590 � 9 578 � 14 582 � 16 585 � 8Pyruvate 569 � 8 582 � 14 558 � 10 580 � 11 560 � 13Succinate 391 � 11 405 � 7 401 � 10 406 � 12 403 � 10

AlcoholsGlycerol 604 � 11 613 � 13 628 � 14 655 � 13 599 � 15Mannitol 630 � 13 623 � 16 625 � 11 634 � 9 622 � 12

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ΔcheAI and ΔcheBI mutants when glucose, sucrose, or xylose was used as the carbonsource, while a less drastic decrease was observed in the presence of other carbonsources (50% for maltose and 25% for tryptone and PYE media). The swimmingbehaviors of the double mutants confirm that the major chemotaxis cluster is the onlyone involved in chemotaxis under the conditions tested here. Both the ΔcheAI ΔcheAIIand the ΔcheBI ΔcheBII mutants phenocopied the ΔcheAI and ΔcheBI single mutantsand formed smaller swim rings than did the WT (Fig. 2). The ΔcheAI ΔcheBI mutant

FIG 2 Motility assays in semisolid agar. (A) Representative images of swim rings obtained after 5 days of incubation in semisolid plates made with M2medium 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 carbonsources. Results are normalized to the WT ring diameter on the same plate type. Bar graphs indicate the mean from five independent replicates, and errorbars 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.

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showed a decreased swim ring similar to those of the ΔcheAI and ΔcheBI singlemutants, confirming that these two genes act in the same pathway to regulatechemotaxis (Fig. 2). Taken together, our results strongly suggest that only the majorchemotaxis cluster is involved in chemotaxis in C. crescentus. It is possible, however,that the alternate chemotaxis cluster is used for chemotaxis under some conditionsother than the ones tested in this study.

It is also worth noting that, under our experimental conditions, the mutants in themajor chemotaxis cluster exhibited a moderately reduced swim ring compared to theWT (25% to 75% reduction depending the tested carbon source) (Fig. 2), while previousworks reported more drastic reductions, similar to a nonmotile mutant (28, 55, 65). Toverify these results, we tested other strains of CheA and CheB mutants described in theliterature as chemotaxis deficient (65, 66); in our hands, all chemotaxis mutants wereable to form larger swim rings than did a nonmotile ΔflgE mutant control, but stillsmaller than did the WT (Fig. S3A). The amplitude of the response was dependent onthe 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 usingNoble agar (0142-01; BD Difco). Interestingly, while M2 plates with no carbon sourceadded did not support growth when made with Noble agar, some growth wasnoticeable using Bacto agar (Fig. S3B), suggesting that Bacto agar contains enoughcarbon traces to be metabolized by C. crescentus.

Both the major and alternate chemotaxis clusters are involved in biofilmregulation. As chemotaxis is involved in surface colonization and biofilm formation indifferent 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 ΔcheAImutant 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 mutantcorresponded 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 certaincarbon sources, as follows: while biofilm formation of the ΔcheAII mutant was notsignificantly different from that of the WT when grown with glucose, sucrose, ortryptone, it dropped to ΔcheAI mutant levels in M2 supplemented with maltose orxylose (Fig. S4). Intriguingly, both the ΔcheBI and ΔcheBII mutants formed more biofilmthan did the WT when grown in defined medium (Fig. S4), suggesting that both CheBproteins are somehow involved in negatively regulating biofilm formation. Mutantphenotypes could be rescued by expressing a copy of the deleted gene on a replicatingplasmid 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, wemonitored their attachment kinetics in the same static system. We focused on PYE andM2 supplemented with glucose (M2G) or xylose (M2X), since the adhesion phenotypesof 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 inbiofilm initiation, with only 30% of the biomass attached compared to the WT strainafter the first 10 h, but this strain eventually caught up and formed as much biofilm asdid the WT at later time points, suggesting that this strain is impaired in early stages ofbiofilm formation only, when cells are grown in PYE (Fig. 3A, left). However, whenglucose was used as the sole carbon source, the ΔcheAI mutant was affected in bothbiofilm 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, ΔcheAIImutant adhesion was similar to that of the WT. Finally, in M2X, when xylose was thesole carbon source, both ΔcheAI and ΔcheAII mutants formed 50% less biofilm than didthe WT (Fig. 3A, right). In all cases, the ΔcheAI ΔcheAII double mutant phenocopied theΔcheAI single mutant (Fig. 3A). As CheAI is involved in chemotaxis (Fig. 2), we concludethat chemotaxis and biofilm formation pathways are interconnected through thiscrucial histidine kinase. In addition, even if CheAII is not involved in chemotaxis (Fig. 2),

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this protein plays a role in biofilm regulation. This regulation is carbon source specificand is not dependent on chemotaxis.

We then assayed CheB mutants (Fig. 3B). In PYE, the amount of biofilm formed overtime 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 complexmedium. However, in M2G and M2X, adhesion was more efficient in both CheB mutantsthan in the WT, with overall 1.5 to 2 times more cells attached to the surface at anygiven time point (Fig. 3B). The amount of biofilm formed by the ΔcheBI ΔcheBII double

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 thewells over time was quantified using crystal violet. Values are given as the average from crystal violet staining of triplicatesamples of at least two independent experiments. The y axis error is represented as the SEM. (A) cheA mutants. (B) cheBmutants. (C) Major cluster mutants. (D) Minor cluster mutants.

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mutant was similar to that of each single mutant. Interestingly, the ΔcheAI ΔcheBI andΔcheAII ΔcheBII double mutants both exhibited a hyperbiofilm phenotype, with anincreased adhesion similar to the cheB single mutants (Fig. 3C and D). This suggests thatCheB acts downstream of CheA in regulating adhesion in C. crescentus.

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

CheA and CheB proteins regulate holdfast production in an antagonistic man-ner. 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 todetermine 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 fluorescentwheat germ agglutinin (WGA), which specifically stains the N-acetylglucosamine resi-dues present in the holdfast (37), and quantified the number of cells with a holdfast byfluorescence microscopy.

As previously reported, the number of cells harboring a holdfast is drastically differentwhen the cells are grown in complex PYE medium than when grown in nutrient-definedM2X (47) or M2G (48) medium. Interestingly, there was also a difference in holdfastformation 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 than10% did in the presence of xylose (Fig. 4A). In all tested media, the number of cellsproducing a holdfast in the ΔcheAI mutant population was reduced by half comparedto the WT (Fig. 4A). In PYE, the number of holdfasts detected in the other testedmutants did not significantly differ from the WT population (Fig. 4A). However, indefined media, both of the �cheB mutant populations produced approximately 20%more holdfasts than did the WT (Fig. 4A). The number of holdfasts present in theΔcheAII 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. 3and S4.

Transcription of the holdfast inhibitor-encoding gene, hfiA, is regulated byCheA and CheB. In C. crescentus, holdfast production is regulated by nutrient avail-ability via the holdfast inhibitor protein HfiA (47). In defined M2G and M2X media, hfiAexpression is upregulated, resulting in a significant decrease in holdfast productioncompared to cells grown in PYE (47, 48). To determine if HfiA was playing a role in thedifferences in holdfast production observed in the present study, we measured theexpression 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 ΔcheAImutant was approximately 50% higher than that in the other tested strains (Fig. 4B). Indefined M2 media, we observed a drastic increase in hfiA transcription compared to inPYE, as previously reported (47–49) (Fig. 4B). There was also an overall increase in hfiAexpression in M2X compared to that in M2G. In defined M2 media, hfiA expression waselevated 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 lowerin 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 testedChe mutants. We first quantified biofilm formation in hfiA che mutants (Fig. 4C). In PYEmedium, the ΔhfiA mutant produced slightly more biofilm than did the WT, as ex-pected. All cheA and cheB single mutants produced similar amounts of biofilm com-pared to the WT strain, and the hfiA che double mutants all phenocopied the ΔhfiA

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FIG 4 Role of the holdfast inhibitor HfiA. (A) Quantification of cells harboring a holdfast in mixedpopulations. Cells were stained using Alexa Fluor 488-WGA and imaged by fluorescence microscopy. Theresults represent the average from three independent replicates (more than 300 cells per replicate), andthe error bars represent the SEM. (B) �-Galactosidase activity of PhfiA-lacZ transcriptional fusions in PYE,M2G, and M2X media supplemented with tetracycline. The results represent the average from 9independent cultures (assayed on 3 different days), and the error bars represent the SEM. Statisticalcomparisons to the WT were calculated using unpaired t tests. (C) Biofilm formation after 24 h ofincubation at 30°C in PYE and M2 media supplemented with glucose or xylose. The results are normalizedto the WT biofilm formation in the given medium. Error bars represent the SEM from three independentreplicates 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.

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mutant in this complex medium. In M2G and M2X defined media, the double mutantsalso phenocopied the ΔhfiA mutant. Based on these results, we conclude that the chegenes and hfiA are acting in the same pathway to regulate holdfast production. Biofilmformation by strains where hfiA is chromosomally inserted at the xylX locus in the chemutants and induced shows a reduction of 80 to 90% compared to the WT emptyvector strain (Fig. S6). This result further suggests that hfiA acts downstream of theholdfast regulation driven by the che proteins. Overall, these observations show thatchemotaxis genes regulate hfiA expression and thereby control holdfast production inresponse to the carbon sources available in the medium.

DISCUSSION

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

Other chemotaxis proteins, specifically, the CheY-like Cle proteins, have been shownto 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 chemotaxisregulation, confirming our observations that only the major chemotaxis cluster prop-erly functions as a regulator of chemotaxis (61). In addition, Cle proteins are involvedin the regulation of holdfast production upon surface contact (61). However, holdfastsynthesis by surface contact stimulation does not occur in defined M2 medium (48),suggesting that the regulation observed in our work is linked to developmentallyprogrammed holdfast production and therefore occurs via a different mechanism. Themultifunctional response regulator MrrA, which is essential for the general stressresponse in C. crescentus, has been recently shown to play a role in chemotaxis andholdfast production via the modulation of hfiA expression (67). It could be interestingto 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 messengermolecule is involved in chemotaxis (61), holdfast production (44, 45), and hfiA expres-sion (48, 49), and it could be involved in the modulation of holdfast synthesis by thechemotaxis proteins.

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

The best-characterized chemotaxis-like operon involved in biofilm formation is the

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Wsp system of Pseudomonas aeruginosa. This system controls EPS production bymodulating cdG levels in response to contact with a surface (73, 74). Briefly, WspR is aCheY homolog and a diguanylate cyclase that acts as the final response regulator of theWsp system (73, 75). In its active form, WspR produces cdG, which in turn activates theproduction 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 increasedphosphorylation of WspR, which negatively regulates the polysaccharides Psl and Pelwhile interfering with the intracellular levels of cdG (73, 75). Future work will determineif holdfast regulation by CheA/CheB in C. crescentus involves a similar mechanism.

In conclusion, we have demonstrated that the two chemotaxis clusters of C. cres-centus have distinct roles. Our data show that while the major cluster is involved in bothchemotaxis and holdfast production, the alternate cluster is a chemotaxis-like systeminvolved in holdfast regulation but not chemotaxis toward the compounds tested.Mutants lacking the kinases CheAI and CheAII are impaired in cell attachment, resultingfrom a defect in holdfast production, while CheBI and CheBII mutants produce moreholdfasts and form more robust biofilms. We also showed that the regulation ofholdfast 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 CheAproteins promote holdfast synthesis, while CheB proteins repress it, by modulating hfiAexpression. Further identification of players in this regulatory pathway and morein-depth exploration of the mechanism by which this occurs may reveal how bacteriarespond to external stimuli to optimize bacterial adhesion and surface colonization invarious environments.

MATERIALS AND METHODSBacterial 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 M2medium (76) supplemented with 0.2% (wt/vol) of a given carbon source (listed in Table 2) or in complexpeptone-yeast extract (PYE) medium (59). When appropriate, 5 �g/ml kanamycin or 1 �g/ml tetracyclinewas 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 �

select cells (Bioline) were used for cloning and were grown in LB medium at 37°C with 25 �g/ml kanamycinor 10 �g/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 toamplify 500-bp fragments from the upstream and downstream regions of the gene to be deleted. Theprimers designed for these in-frame deletions are listed in Table S3. PCR fragments were gel purifiedusing the Zymoclean gel DNA recovery kit (Zymo Research) and then digested by BamHI and XhoI orXhoI and HindIII for upstream or downstream fragments, respectively. Purified digested fragments werethen cloned into the suicide vector pNPTS138 that had been digested by BamHI and HindIII. ThepNPTS138-based constructs were transformed into E. coli Silver � select cells and then introduced intoC. crescentus by electroporation. The two-step recombination was carried out first by selecting integrantson PYE supplemented with kanamycin and second by growing them overnight without selection (into5 ml liquid PYE at 26˚C) and plating the overnight cultures (1 �l) on PYE supplemented with 3% sucroseto select for bacteria that lost the plasmid as part of a second recombination event (77). Then, themutants 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 primerscontaining HindIII (forward primers) and KpnI (downstream primers) restriction sites (Table S2). PCRproducts were gel purified using the Zymoclean gel DNA recovery kit (Zymo Research), digested usingHindIII and KpnI, and ligated into plasmid pMR10 (78), extracted using the Zippy Plasmid prep kit (ZymoResearch), and digested by the same enzymes.

Growth curves and generation time calculations. Bacterial growth in the different media wasmeasured in 3-ml liquid cultures (in 15-ml glass tubes) with shaking at 300 rpm. Overnight cultures werediluted in the same culture medium to an optical density at 600 nm (OD600) of 0.05 and incubated for24 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 singleexponential-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 carbonsource and 0.4% (wt/vol) Noble agar (reference 0142-01; Difco). Cells were stabbed in the soft agar andincubated in a humid chamber at 30°C for 5 days. The diameter of the swimming ring formed by eachtested strain was measured manually.

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Biofilm assays. Biofilm assays in multiwell plates were performed using two different setups thatyield similar results (79), as follows: (i) adhesion to polyvinyl chloride (PVC) microscope coverslips placedvertically in plastic 12-well plates or (ii) adhesion to the inside surface of the wells of untreated plastic24-well plates. Bacteria were grown to mid-log phase (OD600, 0.4 to 0.8) in the chosen medium anddiluted 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 insidesurfaces 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. TheCV from the stained attached biomass was eluted using 10% (vol/vol) acetic acid and was quantified bymeasuring the absorbance at 600 nm (A600). Biofilm formation was normalized to A600/OD600 andexpressed as a ratio of the WT level.

Holdfast quantification using fluorescently labeled WGA lectin. The number of cells harboring aholdfast in mixed populations was quantified by fluorescence microscopy. Holdfasts were detected withAlexa Fluor 488-WGA. Early exponential-phase cultures (OD600, 0.2 to 0.4) were mixed with Alexa Fluor488-WGA (0.5 �g/ml final concentration). One microliter of WGA-stained cells was spotted on a 1.5-mmglass coverslip and covered with an agarose pad (1% SeaKem LE agarose dissolved in dH2O). Holdfastswere 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, andthe Nikon NIS Elements imaging software. The number of individual cells with a holdfast was calculatedmanually from microscopy images in the Nikon NIS Elements imaging software.

�-Galactosidase assays. Strains bearing the transcriptional reporter plasmid of the hfiA genepromoter fused to lacZ (47) were inoculated from freshly grown colonies into 5 ml of a chosenmedium containing 1 �g/ml tetracycline and were then incubated at 30°C overnight. Cultures werethen diluted in the same culture medium to an OD600 of 0.05 and incubated until an OD600 of 0.15to 0.25 was reached. �-Galactosidase activity was measured colorimetrically, as described previously(80). A volume of 200 �l of culture was mixed with 600 �l of Z buffer (60 mM Na2HPO4, 40 mMNaH2PO4, 10 mM KCl, 1 mM MgSO4, 50 mM �-mercaptoethanol), 50 �l of chloroform, and 25 �l of0.1% SDS. Two hundred microliters of the substrate o-nitrophenyl-�-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 �l of 1 M Na2CO3 to raise the pH to 11. A420 was measured,and the Miller units of �-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 theassay. The �-galactosidase activity of WT CB15/plac290 (empty vector control) was used as a blanksample reference.

SUPPLEMENTAL MATERIALSupplemental material for this article may be found at https://doi.org/10.1128/JB

.00071-19.SUPPLEMENTAL FILE 1, PDF file, 0.7 MB.

ACKNOWLEDGMENTSWe 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 themanuscript.

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

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