Azospirillum brasilense Chemotaxis Depends on Two SignalingPathways Regulating Distinct Motility Parameters

Tanmoy Mukherjee,a Dhivya Kumar,a* Nathan Burriss,a Zhihong Xie,a,b Gladys Alexandrea

Department of Biochemistry, Cellular and Molecular Biology, The University of Tennessee, Knoxville, Tennessee, USAa; Yantai Institute of Coastal Zone Research, ChineseAcademy of Sciences, Yantai, Shandong, People’s Republic of Chinab

ABSTRACT

The genomes of most motile bacteria encode two or more chemotaxis (Che) systems, but their functions have been characterizedin only a few model systems. Azospirillum brasilense is a motile soil alphaproteobacterium able to colonize the rhizosphere ofcereals. In response to an attractant, motile A. brasilense cells transiently increase swimming speed and suppress reversals. TheChe1 chemotaxis pathway was previously shown to regulate changes in the swimming speed, but it has a minor role in che-motaxis and root surface colonization. Here, we show that a second chemotaxis system, named Che4, regulates the probability ofswimming reversals and is the major signaling pathway for chemotaxis and wheat root surface colonization. Experimental evi-dence indicates that Che1 and Che4 are functionally linked to coordinate changes in the swimming motility pattern in responseto attractants. The effect of Che1 on swimming speed is shown to enhance the aerotactic response of A. brasilense in gradients,likely providing the cells with a competitive advantage in the rhizosphere. Together, the results illustrate a novel mechanism bywhich motile bacteria utilize two chemotaxis pathways regulating distinct motility parameters to alter movement in gradientsand enhance the chemotactic advantage.

IMPORTANCE

Chemotaxis provides motile bacteria with a competitive advantage in the colonization of diverse niches and is a function en-riched in rhizosphere bacterial communities, with most species possessing at least two chemotaxis systems. Here, we identify themechanism by which cells may derive a significant chemotactic advantage using two chemotaxis pathways that ultimately regu-late distinct motility parameters.

Bacterial chemotaxis provides a competitive advantage by guid-ing motile cells in gradients of chemoeffectors toward envi-

ronments that support growth and metabolism. Chemotaxis con-tributes to the establishment of various associations of bacteriawith eukaryotic hosts (mammals, insects, and plants) and pro-motes virulence, symbiosis, and the establishment of microbialcommunities (1). Bacterial chemotaxis and motility are wide-spread traits encoded in the genomes of bacteria inhabiting di-verse environments, and these functions are specifically enrichedin microorganisms found in soils (2), suggesting that they providea significant competitive advantage in this environment. Consis-tent with these findings, comparative genome analyses of che-motaxis in diverse motile bacteria suggested that most bacteriapossess two chemotaxis systems and soil-dwelling bacteria oftenhave more than two chemotaxis systems (3).

The molecular mechanism of chemotaxis signal transductionhas been deciphered in most detail in the model organism Esche-richia coli, which possesses a single chemotaxis system. In E. coli,the chemotaxis signal transduction pathway consists of mem-brane-bound receptors clustered in dense arrays at the cell poles,where their C-terminal domains associate with cytoplasmic CheAkinase and the CheW scaffolding protein. When stimulated by arepellent, CheA autophosphorylates on a conserved histidine res-idue (H48) using ATP and transfers its phosphate to the aspartateresidue (D57) of CheY (1). Phospho-CheY binding to flagellummotors with a high affinity triggers a switch in the direction offlagellum motor rotation from counterclockwise to clockwise.This event causes a change in the swimming direction of the cell ora tumble (1). A phosphatase, CheZ, assists signal termination by

acting on phospho-CheY (1). In addition, a receptor-specificmethyltransferase, CheR and a receptor-specific methylesterase,CheB, activated by phosphotransfer from phospho-CheA, differ-entially methylate the receptors to reset sensitivity (1). This basicset of chemotaxis proteins comprises a signaling pathway that isgenerally conserved across bacterial species (3). However, thereare notable exceptions to this paradigm, such as the absence ofCheB and CheR in some species, the existence of multiple CheYresponse regulators in others, or the presence of ancillary che-motaxis proteins not found in E. coli (1). Another variation on thistheme is the presence of multiple chemotaxis pathways in thegenomes of most motile bacteria (3). Some of these additionalchemotaxis pathways regulate type IV pilus-dependent motility oralternative cellular functions (ACFs), and they can be identified

Received 8 January 2016 Accepted 4 April 2016

Accepted manuscript posted online 11 April 2016

Citation Mukherjee T, Kumar D, Burriss N, Xie Z, Alexandre G. 2016. Azospirillumbrasilense chemotaxis depends on two signaling pathways regulating distinctmotility parameters. J Bacteriol 198:1764 –1772. doi:10.1128/JB.00020-16.

Editor: I. B. Zhulin, University of Tennessee

Address correspondence to Gladys Alexandre, galexan2@utk.edu.

* Present address: Dhivya Kumar, Department of Molecular Biology andBiophysics, University of Connecticut Health Center, Farmington, Connecticut,USA.

Supplemental material for this article may be found at http://dx.doi.org/10.1128/JB.00020-16.

Copyright © 2016, American Society for Microbiology. All Rights Reserved.

crossmark

1764 jb.asm.org June 2016 Volume 198 Number 12Journal of Bacteriology

on Decem

ber 11, 2020 by guesthttp://jb.asm

.org/D

ownloaded from

due to their unique structure (3). Most chemotaxis pathwaysfound in bacterial genomes are predicted to regulate flagellar mo-tility patterns. The contribution of multiple chemotaxis systems tothe regulation of changes in the swimming direction was demon-strated in some species. For example, in Rhodobacter sphaeroides,two chemotaxis systems together control the probability of stopsin flagellar rotation (4).

Azospirillum brasilense is a motile soil bacterium that inhabitsthe rhizosphere of diverse plant species. A. brasilense cells swimusing a single polar flagellum and change swimming directionwhen the direction of flagellar rotation briefly reverses, causingcells’ movement to be redirected in a different direction (5). Inaddition to regulating the probability of swimming reversals, mo-tile A. brasilense cells navigating an attractant gradient can tran-siently increase their swimming speed (6). The available genomesequence of A. brasilense indicates the presence of four distinctchemotaxis operons, three (che1, che2, and che3) of which are alsopresent in the genomes of all Azospirillum strains sequenced todate as well as in the genome of the closely related organism Rho-dospirillum centenum (7–10), suggesting that they were present inthe last common ancestor of these two genera (Fig. 1). The che-motaxis pathway that regulates transient increases in swimmingspeed in response to attractants has been identified to be Che1 (6).The existence of a distinct pathway for controlling swimming re-versals comes from the observation that inactivation of cheB1 orcheR1 impaired the ability to regulate the probability of swimmingreversals but mutations in either che1, cheA1, or cheY1 did not(11). These data were interpreted to suggest that Che1 function-ally interacts with the unidentified pathway controlling the prob-ability of swimming reversals (6). However, the chemotaxis path-way(s) responsible for controlling changes in the direction offlagellar rotation to trigger reversals is not yet known. The role ofche2 in A. brasilense is unknown, but it is a homolog to the che2operon controlling flagellar biosynthesis in R. centenum (10). In A.brasilense, che2 does not appear to be expressed under standardlaboratory conditions (Z. Xie and G. Alexandre, unpublisheddata). Che3 is a chemotaxis-like ACF pathway that was recentlyimplicated in the control of flocculation in A. brasilense (12).

che4 is present in the genomes of all Azospirillum strains se-quenced to date but absent from the R. centenum genome. In R.centenum, the Che1 chemotaxis system controls all chemotaxisresponses (13); in contrast, its homolog in A. brasilense (Che1)

controls swimming speed and has only a minor role in che-motaxis. The Che4 system is thus the most likely candidate forcontrolling the probability of swimming reversals in this species.Here, we show that Che4 is essential for all chemotaxis responsesin A. brasilense and for competitive wheat root surface coloniza-tion. We demonstrate that the signaling output from Che4 di-rectly modulates the probability of reversals and that signalingfrom Che1 and Che4 is integrated during chemotaxis to producean enhanced response to attractants. These results illustrate anovel mechanism by which motile bacteria utilize two chemotaxispathways regulating distinct motility parameters to alter move-ment in gradients and to increase their chemotactic advantage.

MATERIALS AND METHODSBacterial strains and growth conditions. The bacterial strains used in thisstudy are listed in Table 1. The A. brasilense strains were grown at 28°Cwith shaking (200 rpm). Minimal medium for A. brasilense (MMAB) wasprepared as described previously (14, 15). Cells were induced for nitrogenfixation by first growing them in MMAB, followed by three to four washesof the cell pellet in sterile chemotaxis buffer (10 mM phosphate buffer [pH7.0], 1 mM EDTA) (11) by centrifugation. The pellet was then resus-pended in MMAB lacking any nitrogen source. Nitrogen fixation wasinduced by growth under these conditions overnight at 28°C withoutshaking to ensure low aeration conditions. All culture stocks were rou-tinely maintained on tryptone-yeast (TY) medium (per liter, 10 g Bactotryptone, 5 g yeast extract) or MMAB solidified with 1.5% (wt/vol) agar.For A. brasilense, the following antibiotics were added at the indicatedfinal concentrations: ampicillin at 200 �g/ml, chloramphenicol at 20 �g/ml, kanamycin at 30 �g/ml, gentamicin at 20 �g/ml, and tetracycline at 10�g/ml.

Mutagenesis. To construct a strain from which the cheA4 gene wasdeleted, an 843-bp upstream fragment including 126 bp of cheA4 and adownstream 825-bp sequence including 232 bp at the 3= end of cheA4 werePCR amplified using primers cheA4Up-F, cheA4Up-R, cheA4Dwn-F, andcheA4Dwn-R (see Table S1 in the supplemental material). These primerswere engineered to include 5= XbaI (cheA4Up-F) and 3= HindIII(cheA4Dwn-R) restriction sites for cloning into pUC19 as well as BamHIsites (present at the 5= end of primer cheA4Up-R and at the 3= end ofprimer cheA4Dwn-F) to permit subsequent insertion of a gentamicin re-sistance (Gmr) cassette isolated from p34S-Gm by BamHI restriction di-gestion (Table 1). After verification of the sequence by sequencing, the�cheA4::Gm region present on the pUC19 vector was isolated by restric-tion digestion with XbaI and HindIII and inserted into the suicide vectorpSUP202 (Table 1) that had been digested with the same enzymes, yield-ing pSUP�cheA4::Gm. The pSUP�cheA4::Gm vector was transformedinto E. coli S17-1 for biparental mating with strains A. brasilense Sp7 andits �cheA1 mutant derivative for allelic exchange, as previously described(15). For constructing a mutant lacking cheY4, an upstream fragmentencompassing the first 6 bp of cheY4 and an additional 577 bp of upstreamDNA sequence and a downstream fragment including the last base pair ofcheY4 and an additional 584 bp downstream of the stop codon were am-plified using the primer pair cheY4Up-F and cheY4Up-R and the primerpair cheY4Dwn-F and cheY4Dwn-R (see Table S1 in the supplementalmaterial), which were designed to include restriction sites to facilitatecloning into the EcoRI- and HindIII-digested pUC19 vector. The frag-ments were also engineered to include a BamHI site for insertion of achloramphenicol cassette isolated from a p34S-Cm BamHI-digested vec-tor. The cheY4 deletion insertion construct in pUC19 was verified by se-quencing. The sequence-verified fragment was isolated by digestion withEcoRI and HindIII and cloned into the pKGmobGII suicide vector toyield pKG�cheY4::Cm (Table 1). The latter was transformed into E. coliS17-1 cells and transferred into A. brasilense Sp7 or its �cheY1 mutantderivative by biparental mating for allelic exchange, as described above.PCR was used to verify that the strains carried the appropriate deletions.

cheY1 cheB1 cheR1

cheW2 cheB2 orf cheR2 cheY2 cheA2 orf

mcp cheA3 cheB3 RR HK

cheW4 mcp mcpcheB4cheD4cheR4

Che1

Che4

Che2

Che3

cheA1 cheW1

cheY3 cheC mcp

cheW3a cheR3 cheW3b

cheA4cheScheY4

FIG 1 Chemotaxis gene clusters within the A. brasilense genome. Boxes rep-resent open reading frames and are drawn to scale. The chemotaxis geneswithin each cluster were either previously characterized or identified by ho-mology searches, as detailed in the text. orf, open reading frame; RR, responseregulator gene; HK, histidine kinase gene; mcp, gene for methyl-acceptingchemotaxis protein.

Chemotaxis Signal Transduction in Azospirillum

June 2016 Volume 198 Number 12 jb.asm.org 1765Journal of Bacteriology

on Decem

ber 11, 2020 by guesthttp://jb.asm

.org/D

ownloaded from

To construct a strain from which most of the che4 sequence was deleted,we first used PCR amplification of a 698-bp fragment encompassing thefirst gene of the che4 cluster (Fig. 1) and an upstream DNA sequence usingprimers che4Up-F and che4Up-R (see Table S1 in the supplemental ma-terial) and a 722-bp fragment overlapping the last gene of che4 (Fig. 1) andincluding an additional downstream DNA sequence using primersche4Dwn-F and che4Dwn-R (see Table S1 in the supplemental material).The primers che4Up-R and che4Dwn-F were designed to include 12 bp ofoverlapping DNA sequence that was used in a second round of splicing byoverhang extension PCR (16) to produce a 1,420-bp fusion product be-tween the upstream and downstream PCR fragments. The 1,420-bp prod-uct was restriction digested with PstI and XbaI, which were engineered inthe che4Up-F and the che4Dwn-R primers, respectively (see Table S1 inthe supplemental material), and cloned into pUC19. The fusion constructcloned in pUC19 was further digested with SalI in order to insert a SalI-digested gentamicin resistance cassette isolated from the p34S-Gm vector(Table 1) and yield a �che4::Gm construct cloned in pUC19. After verifi-cation by sequencing, the �che4::Gm region was isolated as a PstI andXbaI fragment which was then cloned into the suicide vectorpSUPPOL2SCA (17). The suicide vector carrying the che4 deletion inser-tion construct was transformed into E. coli strain S17-1, followed by allelicexchange after transfer to A. brasilense wild-type strain Sp7 and its �che1mutant derivative by biparental mating, as described above. Mutants car-rying the correct deletion insertions were identified by PCR.

Complementation and site-specific mutations of cheA4 and cheY4genes. Functional complementation of the mutant phenotypes was car-ried out by expressing the parental genes or mutated alleles in trans fromthe broad-host-range pRK415 vector (Table 1). The genes were cloned

in-frame and downstream of the plasmid-borne lac promoter with engi-neered restriction sites and a ribosome binding site to ensure expression(Table 1) (6). Each gene was amplified from the genomic DNA of thewild-type strain, using a set of forward and reverse primers that are listedin Table S1 in the supplemental material. The genes coding for kinase-inactive CheA4 with the H54Q substitution (CheA4H54Q) and the inactiveCheY4 with the D57N substitution (CheY4D57N) were synthesized byGenScript (Piscataway, NJ) and cloned into pUC57 between EcoRI andHindIII sites. The cheA4 and cheY4 inserts were subcloned into pCR2.1 byTOPO cloning according to the manufacturer’s instructions (Invitrogen,CA) for verification by sequencing, followed by digestion with EcoRI andHindIII to isolate the sequence-verified genes. The isolated DNA frag-ments were cloned into the pRK415 vector. Recombinant plasmids wereisolated from transformed E. coli TOP10 cells, prior to being transferredinto E. coli S17-1 for biparental mating with A. brasilense recipient strains,as described above. An empty pRK415 vector was transferred to A.brasilense Sp7 and its mutant derivatives for use as controls.

Behavioral assays. For the capillary assay for aerotaxis, cells weregrown to an optical density at 600 nm (OD600) of 0.4 to 0.6 (exponentialphase of growth) in MMAB supplemented with malate (10 mM) andammonium (18.7 mM). Cultures were adjusted to an equivalent numberof cells (estimated via OD600 measurements) by dilution in chemotaxisbuffer. The cells were gently washed three times with chemotaxis buffer bylow-speed centrifugation and resuspended in 100 �l MMAB containingmalate (10 mM). All cells remained motile under these conditions. Cellswere transferred to an optically flat capillary tube (Vitro Dynamics, Inc.,Rockaway, NJ) by immersing a capillary tube into the cell suspension.Aerotaxis was visualized under a light microscope as the formation of a

TABLE 1 Strains and plasmids used in this study

Strain or plasmid Genotype or description Reference or source

StrainsAzospirillum brasilense

Wild type Wild-type strain Sp7 ATCC 29145AB101 �cheA1::gusA-Km (Kmr) 24AB102 �cheY1::Km (Kmr) 24AB103 �(cheA1-cheR1)::Cm (Cmr) 24AB401 �cheA4::Gm (Gmr) This workAB402 �cheY4::Cm (Cmr) This workAB403 �(cheX4-cheD4)::Gm (Gmr) �che4 This workAB141 �cheA1 �cheA4::gusA-Km-Gm (Kmr Gmr) This workAB142 �cheY1 �cheY4::Km-Cm (Kmr Cmr) This workAB143 �che1 �che4::Cm-Gm (Cmr Gmr) This work

Escherichia coliTOP10 General cloning strain InvitrogenS17-1 thi endA recA hsdR strain with RP4-2Tc::Mu-Km::Tn7 integrated in

chromosome42

PlasmidspRK415 Cloning vector (Tcr) 43pRKCheA4 pRK415 containing cheA4 This workpRKCheA4H54Q pRK415 containing cheA4 with a mutation replacing residue H54 with Q This workpRKCheY4 pRK415 containing cheY4 This workpRKCheY4D57N pRK415 containing cheY4 with a mutation replacing residue D57 with N This workpSUP202 Suicide vector; pBR325 mob Tcr Ampr Cmr 42pSUP�cheA4::Gm pSUP202 with �cheA4::Gm; Gmr This workpKGmobGII Kmr; mobilizable suicide vectorpKG�cheY4::Cm pKGmobGII with �cheY4::Cm; Cmr

p34S-Gm ori ColE1 Ampr Gmr 44p34S-Cm ori ColE1 Ampr Cmr 44pUC19 General cloning vector; Ampr 45pSUPPOL2SCA Derivative of pSUP202; oriT of RP4; Tcr 17

Mukherjee et al.

1766 jb.asm.org June 2016 Volume 198 Number 12Journal of Bacteriology

on Decem

ber 11, 2020 by guesthttp://jb.asm

.org/D

ownloaded from

stable band of motile cells at a distance from the air-liquid interface (themeniscus). An aerotaxis band typically forms within 2 to 3 min and isstable for a minimum of 25 min under these conditions (18). The assaywas performed in triplicate using independent cultures. Images were cap-tured using the 4� objective of a phase-contrast Nikon E200 microscopevia a C-mounted Nikon Coolpix digital camera.

For chemotaxis in a spatial gradient, soft agar plates were preparedusing MMAB solidified with 0.3% agar and supplemented with malate (10mM) and ammonium chloride (18.7 mM). Cells were grown as indicatedabove for aerotaxis, and the cell suspensions were adjusted to an equiva-lent number of cells, estimated by OD600 measurements, prior to beinginoculated in the center of the soft agar plates. The inoculated plates wereincubated at 28°C for 48 h before being photographed. The experimentwas performed in triplicate for each strain.

Computerized motion tracking of free-swimming cells. Digitalmovies of free-swimming cells were captured using the 40� objective of aphase-contrast Nikon Eclipse E200 microscope fitted with a Sony Hyper-HAD monochrome camera at a rate of 30 frames per second. The videoswere converted to a digital format using IC Capture2 software (The Im-aging Source, Charlotte, NC) before being analyzed using CellTrak (ver-sion 1.5) software (Motion Analysis, Santa Rosa, CA). Changes in swim-ming parameters (velocity and reversal frequency) were determined for aminimum of 75 cells recorded from at least two different fields of view andthree independent cultures. Velocity is expressed as micrometers per sec-ond, and reversal frequency is expressed as the number of swimmingreversals per second and per cell.

Plant inoculation. Wheat (Triticum aestivum) seeds were surface ster-ilized and germinated for 3 days in the dark followed by an additional dayin the light, as previously described (19). Fifty milliliters of Fahraeus me-dium (CaCl2, 100 mg liter�1; MgSO4·7H2O, 120 mg liter�1; KH2PO4, 100mg liter�1; Na2HPO4, 150 mg liter�1; ferric citrate, 5 mg liter�1) solidifiedwith 0.6% agar was placed in a 6.5-cm-diameter growth chamber. Fourplantlets were placed at an equal distance from each other and from thecenter of the growth chamber. Strains to be tested were inoculated at thecenter of the chamber. Eight chambers (with four plants each) were pre-pared per trial, yielding a minimum of 8 replicates. For inoculation,strains were grown in MMAB supplemented with malate (10 mM) andammonium chloride (18.7 mM) to an OD600 of 0.5. Two milliliters of theculture was pelleted, washed 3 times with sterile 0.8% KCl by centrifuga-tion, and resuspended in 400 �l of sterile 0.8% KCl. Twenty microliters ofthis suspension was inoculated into the center of the growth chamber. Thenumber of cells in the inoculum was determined by counting the numbersof CFU. For competitive root colonization assays, the strains to be com-pared were prepared as indicated above and mixed in a 1:1 ratio prior toinoculation into the chambers. The chambers were incubated at 25°C for24 h, after which the plants were removed and the roots were excised. Theroots of the four plantlets from a single growth chamber were combinedand homogenized in 5 ml 0.8% KCl, followed by serial dilutions andplating on selective medium to determine the numbers of CFU. Rootcolonization was calculated as a colonization index for the plants inocu-lated with a single strain and as a competitive index for the plants inocu-lated with two strains. The colonization index for an individual strain wasthe log10(strain output/strain input), where the number of CFU extractedfrom roots after incubation was normalized to the number of CFU mea-sured in the inoculated input. The competitive index was a ratio calculatedas follows: log10[(mutant output/wild-type output)/(mutant input/wild-type input)].

Identification of chemotaxis proteins in the complete genome se-quence of A. brasilense. The che1 operon organization was previouslydescribed (14). To identify other chemotaxis proteins, we used the se-quences of the chemotaxis homologs reported in the genome descriptionof A. brasilense Sp245 (10) as queries to search the genome of A. brasilenseSp7 (BioProject accession number PRJNA293508) using the BLASTserver (20).

Statistical analysis. For comparing the wild-type and mutant pheno-types (swimming speed, swimming reversals, and plant colonization), wedetermined average values from at least three independent experimentsperformed in duplicate and performed one-way analysis of variance (al-pha level, 0.05), followed by pairwise two-sample t tests assuming equalvariances (alpha level, 0.05) using Prism (version 6) software (GraphPadSoftware Inc., San Diego, CA).

RESULTSche4 mutants lack chemotaxis and aerotaxis. Genome sequenceanalysis predicted that Che4 is the major chemotaxis system in A.brasilense. To test this hypothesis, we constructed strains lackingcheA4 (�cheA4; strain AB401), cheY4 (�cheY4; strain AB402), orthe entire che4 cluster (�che4; strain AB403) (Table 1) and testedtheir chemotaxis and aerotaxis abilities using spatial gradient as-says (Fig. 2). All mutants were motile and grew at similar rates invarious media and under various incubation conditions (data notshown). While the �cheA4 mutant was null for chemotaxis and

*Sp7 ΔcheA4

ΔcheY4 Δche4

ΔcheA4

ΔcheY4

Δche4

Direction of airdiffusion

ΔcheY4(pRK415)

Sp7(pRK415)

ΔcheA4(pRK415)

ΔcheA4(pRKCheA4)

ΔcheA4(pRKCheA4 )H54Q

ΔcheY4(pRKCheY4)

ΔcheY4(pRKCheY4 )D57N

Sp7

ΔcheA4

ΔcheY4

Δche4

Sp7(pRK415)

A B

C D* *

FIG 2 Taxis behaviors of A. brasilense and its che4 mutant derivatives. (A)Chemotaxis in the soft agar plate assay containing malate (10 mM) and am-monium chloride (18.7 mM) as the carbon and nitrogen sources, respectively.The strains tested are indicated at the top of each plate. The pictures were takenafter 48 h of incubation at 28°C. Representative images from at least five dif-ferent assays are shown. (B) Aerotaxis in the spatial gradient assay. The airgradient is established by diffusion, in the direction indicated by the arrow,into capillary tubes filled with a suspension of motile cells. The images weretaken 5 min after placement of the cells in the capillary tubes. (C and D)Functional complementation of the aerotaxis defects of the �cheA4 mutant(C) and of the �cheY4 mutant (D) in the capillary assay for aerotaxis. Thephotographs were taken 10 min after placement of the suspension of motilecells in the capillary tubes. The plasmids carried by the strains are derivatives ofthe broad-host-range plasmid pRK415. The aerotaxis defect of the �cheA4mutant strain is rescued by expression of wild-type CheA4 but not CheA4H54Q

(C). Similarly, the aerotaxis defect of the �cheY4 mutant strain is rescued byexpression of a wild-type CheY4 but not CheY4D57N (D). In panels B, C, and D,the number of cells was equivalent in all tubes, and all strains were motile. *,formation of a stable band of motile cells.

Chemotaxis Signal Transduction in Azospirillum

June 2016 Volume 198 Number 12 jb.asm.org 1767Journal of Bacteriology

on Decem

ber 11, 2020 by guesthttp://jb.asm

.org/D

ownloaded from

aerotaxis, the �cheY4 and �che4 mutants were null for aerotaxisbut displayed residual chemotaxis in the soft agar assay (Fig. 2Aand B). Similar patterns were observed when other carbon sourceswere used in the soft agar assay (data not shown). The differentphenotypes of some of these mutants for chemotaxis versus aero-taxis were intriguing. The discrepancy between the behavior of the�cheY4 and �che4 mutants in the aerotaxis versus chemotaxisassay could be due to the different incubation times and condi-tions under which these assays were conducted. The aerotaxis as-say was performed using a suspension of free-swimming cellsplaced into a capillary tube. The aerotactic band typically formswithin 2 to 3 min. The chemotaxis rings formed in soft agar plateswere observed after at least 24 to 48 h of growth. One possibility isthat the rings observed in the soft agar plates were the result ofpseudotaxis, which is a form of translocation through the agar thatdoes not result from chemotaxis signaling; these have been re-ported under similar conditions in several bacterial species (21–23). The aerotaxis defects of the �cheA4 and �cheY4 mutantscould be complemented by expressing a parental gene from abroad-host-range plasmid but not by expressing a variant allele ofcheA4 or cheY4 carrying a single mutation on the predicted phos-phorylatable histidine (H54Q in CheA4) and aspartate (D57N inCheY4) residues, respectively (Fig. 2C and D). These data indicatethat conserved phosphorylatable residues on CheY4 and CheA4must be present for signaling to occur, suggesting that a phos-phorylation cascade between CheA4 and CheY4 triggers changesin the direction of flagellar rotation to cause swimming reversals.Together, these results identify CheA4 to be the major histidinekinase mediating aerotaxis and chemotaxis in A. brasilense, andthey further suggest that signaling from Che4 plays a major role incontrolling the changes in the direction of flagellar rotation duringswimming reversals.

The signaling output from Che4 is the control of the swim-ming reversal frequency. As indicated above, the formation ofchemotaxis rings in the soft agar assay and the formation of aero-taxis bands in the capillary assay are not identical behaviors. Theultimate signaling output of a bacterial chemotaxis pathway is thecontrol of the swimming motility pattern by affecting the proba-bility of reversals. In other words, mutants unable to chemotax areexpected to either constantly run or constantly reverse swimmingdirection. We assessed the swimming motility patterns of the�che4, the �cheA4, and the �cheY4 mutants under steady-stateconditions (Fig. 3A). The wild-type strain swam with long runsinterrupted by instances of reversals that occurred with an averageprobability of about 0.5 reversal/s (Fig. 3B). In contrast to the wildtype, mutants lacking CheA4 swam in straight runs and did notreverse (Fig. 3A and B). The �cheY4 mutant had a phenotypesurprisingly different from that of the �cheA4 mutant, in that italso swam with fewer instances of reversals than the wild-typestrain (Fig. 3A and B), but it clearly was still able to reverse swim-ming direction (Fig. 3A). The strain lacking the che4 cluster(�che4) displayed a frequency of reversals that was not signifi-cantly different from that of the wild-type strain (0.6 reversal/s)(Fig. 3A and B). Analysis of free-swimming cells of the �che4

ΔcheY1ΔcheY4

ΔcheA1ΔcheA4Sp7

Δche1Δche4

C

Swim

min

g re

vers

al fr

eque

ncy

(r

ever

sal/s

)

ΔcheA

1Δch

eA4

Sp7

ΔcheA

4

ΔcheY

4Δc

he4

ΔcheY

1Δch

eY4

Δche1Δ

che4

Sp7 ΔcheA4

ΔcheY4 Δche4

A

B

FIG 3 Swimming behavior of the A. brasilense wild type and mutant deriva-tives lacking che4 genes or lacking a combination of che4 and che1 genes. (A)Tracks of free-swimming cells of A. brasilense and its che4 mutant derivatives.The tracks were obtained from digital recordings and computerized motionanalysis using CellTrak software. Representative tracks are shown. Arrows,instances of swimming reversals. (B) Reversals were determined by comput-erized motion analysis from tracks of at least 75 free-swimming cells fromthree independent cultures. The results shown are the average numbers ofswimming reversals per second for each strain analyzed with standard devia-tions. *, statistically significantly different values compared to the value for thewild-type strain (t test; P � 0.05). The probabilities of reversal of the �cheA1,�cheY1, and �che1 mutant strains under similar conditions were previouslypublished (24) and corresponded to 0.41 � 0.1 reversal/s, 0.37 � 0.04 rever-sal/s, and 0.46 � 0.05 reversal/s, respectively. (C) Tracks of free-swimming

cells of A. brasilense and its derivatives lacking both che1 and che4 genes. Thetracks were obtained from digital recordings and computerized motion anal-ysis (CellTrak). Representative tracks are shown. Arrows, instances of swim-ming reversals.

Mukherjee et al.

1768 jb.asm.org June 2016 Volume 198 Number 12Journal of Bacteriology

on Decem

ber 11, 2020 by guesthttp://jb.asm

.org/D

ownloaded from

strain also revealed a significantly erratic (jiggly) motility pattern(Fig. 3A). This analysis thus confirms that CheA4 is essential forthe cells to reverse swimming direction. The lack of CheY4 orChe4 severely impaired but did not eliminate the ability of cells toreverse, while a lack of CheA4 completely abolished it, indicatingthat CheY4 (and, thus, Che4) has a major but not unique role inthis behavior.

Che1 and Che4 together contribute to regulating the swim-ming pattern. The distinct swimming pattern of the �cheY4 and�che4 strains compared to that of the �cheA4 strain was unex-pected, since all three strains should display the same phenotype ifthe signaling output of the Che4 pathway is the control of swim-ming reversals. Che1 controls changes in swimming speed duringchemotaxis, and previous data have suggested that signaling fromChe1 and from the pathway affecting swimming reversals func-tionally interacted (6, 24). Therefore, we hypothesized that Che1may be responsible for the residual swimming reversal ability ofthe �cheY4 and the �che4 mutants. To test this possibility, weconstructed strains lacking both che1 and che4 (�che1 �che4;strain AB143), both cheA1 and cheA4 (�cheA1 �cheA4; strainAB141), and both cheY1 and cheY4 (�cheY1 �cheY4; strainAB142) (Table 1) and analyzed their swimming patterns (Fig. 3C).We found that the mutation of cheY1 and che1 in the �cheY4 and�che4 mutant backgrounds yielded cells unable to reverse swim-ming direction, similar to the motility pattern observed for the�cheA4 strain or, as expected, for the �cheA1 �cheA4 strain (Fig.3B and C). The disabling of Che1 in the �che4 mutant (strainAB143) background also abolished the erratic swimming behaviorof the �che4 strain, suggesting that signaling from Che1 causedthis behavior (Fig. 3B and C). The �che1 �che4 mutant strain and,to a lesser extent, the �cheA1 �cheA4 strain also persisted andswam in circles close to the surface of the coverslip (Fig. 3C). Thisbehavior has been associated with smooth swimming close to sur-

faces in E. coli (25, 26) and is thus consistent with the lack ofswimming reversals in these strains. However, we do not knowwhy this behavior is more preeminent in these two mutants butnot in the �cheY1 �cheY4 strain. Since Che1 affects the swimmingspeed, we also analyzed this motility parameter for mutants lack-ing che4 genes alone or in combination with mutations in che1genes. We found that only the �cheA4 and the �che4 mutants hada small, but significant, reduction in steady-state swimming speedcompared to that of the wild-type strain (Fig. 4). This phenotypewas absent in the �cheA1 �cheA4 and �che1 �che4 strains, impli-cating Che1 in the reduced swimming speed. These results suggestthat it is the combination of reduced swimming speed and main-tenance of the ability to reverse that caused the erratic swimmingpattern of the �che4 mutant strain (Fig. 3A). Together, the dataconfirm the role of Che1 and Che4 in regulating the motility pat-terns of swimming cells and in chemotaxis. They also providefurther support to the hypothesis of a functional signaling inter-action between Che1 and Che4.

The aerotactic advantage of coordinated regulation of swim-ming speed and reversals. We wondered what could be the ad-vantage for A. brasilense cells in using two distinct chemotaxissignaling systems regulating two different parameters of swim-ming motility to modulate taxis responses. Che1, via signalingthrough CheY1 (6), regulates transient increases in swimmingspeed. Che4, via signaling through CheY4, controls reversals, asshown here. We hypothesized that a transient increase in theswimming speed that would accompany a suppression of swim-ming reversals during the response to an attractant would enhancethe net movement of cells in this gradient. To test this hypothesis,we compared aerotaxis in a spatial gradient assay of the wild-typestrain with that of its �cheY1 and �cheY4 mutant derivatives in atime course experiment (Fig. 5). As expected, the �cheY4 straindid not form an aerotactic band, although the cells were fullymotile. The wild-type strain started forming an aerotactic bandafter 90 s, and a stable aerotactic band was formed by 150 s. Theinitiation of the aerotactic band took almost twice as much timefor the �cheY1 strain, which formed a stable aerotactic band at 300s postinoculation in the capillary tubes. This delay in the forma-tion of the aerotactic band by the �cheY1 cells relative to the timeof formation for the wild-type strain was consistently reproduc-ible. These results suggest that the control of swimming speed byChe1 enhances the response of A. brasilense cells to a gradient of amajor attractant, oxygen.

Che4 controls plant association. The major role of Che4 incontrolling chemotaxis and the swimming pattern suggested thatit should also play a significant role in the ability of cells to colonizethe roots of cereals, such as wheat, a common host plant for A.

Swim

min

g sp

eed

(µm

/s)

ΔcheA

1Δch

eA4

Δche4

ΔcheY

1Δch

eY4

Δche1Δ

che4

ΔcheY

4

ΔcheA

4Sp7

FIG 4 Swimming speed of free-swimming cells of the A. brasilense wild typeand its mutant derivatives lacking che4 genes or lacking a combination of theche4 and che1 genes. The values shown are average swimming speeds, deter-mined by motion tracking of at least 75 cells from three independent cultures,with standard deviations. *, statistically significantly different values comparedto the value for the wild-type strain (t test; P � 0.05). The swimming speeds ofthe �cheA1, �cheY1, and �che1 mutant strains under similar conditions werepreviously published (35) and corresponded to 22 � 1 �m/s, 31 � 3 �m/s, and28 � 2 �m/s, respectively.

00

Sp7

ΔcheY1

ΔcheY4

90 sec 150 sec 300 sec

FIG 5 Time course of aerotactic band formation in the A. brasilense wild typeand its �cheY1 and �cheY4 mutant derivatives. Cell suspensions adjusted toequivalent numbers of cells were placed in capillary tubes, and recording (thebeginning of which is shown at the upper left of each panel) was started im-mediately (time zero). All cells were motile throughout the experiments. Rep-resentative images are shown.

Chemotaxis Signal Transduction in Azospirillum

June 2016 Volume 198 Number 12 jb.asm.org 1769Journal of Bacteriology

on Decem

ber 11, 2020 by guesthttp://jb.asm

.org/D

ownloaded from

brasilense. When inoculated alone onto sterile wheat plants, thewild type and its �che4 mutant derivative were able to colonizesterile wheat roots (Fig. 6A), but the mutants did so at a relativelyreduced level compared to that for the wild type. Given that che-motaxis provides bacteria with a competitive advantage, we hy-pothesized that the colonization defects of the mutants wouldbecome apparent in competition experiments against the wild-type strain and thus compared the competitive index of the �che4mutant with that of the wild-type strain when they were inocu-lated at a 1:1 ratio (Fig. 6B). The competition experiments showedthat a lack of che4 significantly impaired the competitive ability ofthe strains to colonize the wheat root surfaces, confirming themajor role of this pathway in the lifestyle of A. brasilense in therhizosphere.

DISCUSSION

Experimental evidence shows that Che4 signaling controls swim-ming reversals in A. brasilense and that it is the major pathway forall taxis responses in this species. As expected from its major func-tion in regulating chemotaxis responses, Che4 is essential for com-petitive root surface colonization in A. brasilense. The A. brasilenseChe4 pathway is orthologous to the major pathway controlling allchemotaxis responses in Sinorhizobium meliloti (27, 28) and Rhi-zobium leguminosarum (23), where this chemotaxis system is alsoessential for plant root colonization. che4 is present in the ge-nomes of all Azospirillum strains sequenced to date (7–9) but ab-sent from the phylogenetically related aquatic organism R. cente-

num (10). In R. centenum, the Che1 chemotaxis system controls allchemotaxis responses (13), while its homolog in A. brasilense(Che1) controls swimming speed (6). Furthermore, mutations inA. brasilense che1 did not affect the ability to colonize the surface ofwheat roots, indicating that this pathway does not contribute tothe rhizosphere lifestyle of this organism (19). Lateral gene trans-fer (LGT) is considered a major driving force in the evolution ofprokaryotes, including the acquisition of functions for adaptationto new niches (29). The A. brasilense Che4 pathway was previouslyidentified to be one of the functions acquired by LGT by the an-cestors of Azospirillum spp. that were proposed to have contrib-uted to the adaptation to the rhizosphere (10). The role for Che4in root surface colonization characterized here is thus consistentwith this hypothesis. Che4 is a representative of the F7 class ofchemotaxis systems (3) that was shown to be enriched in rhizo-sphere bacterial communities relative to those found in soil (2).The F7 chemotaxis systems, such as A. brasilense Che4, may thusbe advantageous in this environment. However, the specific com-petitive advantage(s) that signaling via such a chemotaxis pathwayprovides remains to be identified.

The acquisition of Che4 by LGT in the ancestor of Azospirillumspp. did not cause Che1 to lose its function in chemotaxis. Indeed,our results indicate that both Che1 and Che4 have a role in theregulation of the swimming pattern of motile A. brasilense cellsand thus chemotaxis. However, the contribution of Che1 andChe4 to this behavior is significantly different. As shown previ-ously (6) and in the present study, Che1 controls transient in-creases in swimming speed, while Che4 controls the probability ofswimming reversals and plays a major role in all taxis responses.Our data indicate that signaling via Che4 controls most swimmingreversals, with Che1 modulating this behavior via direct effects onspeed. This is in contrast to the role of two chemotaxis pathways inR. sphaeroides, a bacterial species that has been extensively studiedin this respect (30). Two chemotaxis systems, named CheOp2 andCheOp3, are implicated in the control of chemotaxis in motile R.sphaeroides, and the signaling output from both chemotaxis path-ways controls stops in the rotation of flagellar motors (31, 32).Consistent with a single signaling output, inactivation of eitherCheOp2 or CheOp3 abolishes chemotaxis in R. sphaeroides. This isbecause both signaling pathways ultimately affect the activity of acommon set of CheY response regulators that bind the flagellarmotors to stop their rotation (33, 34). This is in contrast to theresults obtained here for A. brasilense that suggest that the signal-ing output from both Che1 and Che4, likely mediated via CheY1(6) and CheY4, respectively, alters different parameters of flagellarmotor activity. The control of chemotaxis by two Che pathways inA. brasilense thus illustrates a distinct strategy by which the signal-ing input from multiple Che pathways regulates chemotaxis.

Several lines of experimental evidence also support the hypoth-esis of a signal integration via a functional interaction betweenChe1 and Che4. First, inactivation of cheA4 completely sup-pressed reversals, and it also caused the cells to swim at a reducedspeed compared to that of cells of the wild-type strain (24, 35).Second, strains lacking the CheY4 or Che4 function were signifi-cantly impaired but not null for chemotaxis unless cheY1 or che1,respectively, was also deleted. The discrepancy between the swim-ming speed phenotype of the �cheA4 and �cheY4 strains indi-rectly supports the hypothesis of a functional interaction betweenChe1 and Che4 because a �cheA1 strain, but not a �cheY1 strain,also swam at a slower speed than the wild-type A. brasilense strain

che4:Sp7-1

0

1

2

3

4

Com

petit

ive

Inde

x

Sp7 che4-1

0

1

2

3

4

Col

oniz

atio

nIn

dexA

B

FIG 6 Role of Che4 in the A. brasilense root surface colonization of wheat. (A)The colonization index was determined from A. brasilense and its �che4 mu-tant derivative inoculated alone onto sterile wheat plantlets and was calculatedas log10(strain output/strain input), where the number of CFU extracted fromroots after incubation was normalized to the number of CFU measured in theinoculated input. Dots, colonization indexes for a set of four plants (onegrowth chamber) inoculated with the strain indicated; horizontal bars, means;vertical lines, standard deviations; broken line, a colonization index of 1, indi-cating that all inoculated cells were recovered from the plant root surface. Theindex was �1 for Sp7, likely because of growth on the roots. The colonizationindexes of the wild type and the �che4 mutant were statistically significantlydifferent (t test; P � 0.0001). (B) The competitive index was determined as theratio of the mutant to the wild type recovered from roots after being inoculatedat a 1:1 ratio (mutant/wild type) and was calculated as log10[(mutant output/wild-type output)/(mutant input/wild-type input)]. Dots, competitive indexfor four plants (one growth chamber) inoculated with the strains indicated ata 1:1 ratio; horizontal bar, the mean; broken line, a competitive index of 1,corresponding to strains equally competitive for root surface colonization.

Mukherjee et al.

1770 jb.asm.org June 2016 Volume 198 Number 12Journal of Bacteriology

on Decem

ber 11, 2020 by guesthttp://jb.asm

.org/D

ownloaded from

(6). The lack of observable swimming speed defects in strains lack-ing CheY1 or CheY4 could result from either a functional redun-dancy between CheY1, CheY4, and any of the 5 other CheY ho-mologs encoded in the genome (10) or some other unidentifiedinteractions between the Che1 and Che4 proteins. The mecha-nism by which signaling from Che1 and Che4 is integrated toaffect swimming speed and reversals remains to be elucidated, butprevious data suggested that chemotaxis receptors and adaptationproteins may be involved in this process (11, 35). This was alsosuggested for the role of Che1 and Che2 in Rhizobium legumino-sarum bv. viciae chemotaxis (23) and was demonstrated to be anexisting mechanism in R. sphaeroides (31, 32).

Increased swimming speed during chemotaxis provides an ad-vantage for marine bacteria in the exploitation of transient sourcesof nutrients in the oceans (36). Like the ocean, the soil is a hetero-geneous environment characterized by a plethora of nutrient gra-dients that change rapidly in space and time (37). In contrast tothe ocean, the soil is less limiting in many essential nutrients, andas a result, it harbors abundant and diverse microbial communi-ties (38). Oxygen may be particularly limiting in the rhizosphere,where microbial cell densities and microbial activities are thegreatest (39, 40), making oxygen a potential modulator of micro-bial activities and competition in this environment. Consistentwith this hypothesis, a major chemotaxis receptor for sensing ox-ygen and other metabolism-related parameters is critical for theability of A. brasilense to colonize the wheat root surface (41). Theresults obtained here suggest that A. brasilense derives an aerotac-tic advantage from increased swimming speed, which is expectedto be significant in the competitive environment of the rhizo-sphere. The additive roles of two chemotaxis systems controllingdistinct motility parameters, illustrated here by the A. brasilenseChe1 and Che4 systems, suggest a potential benefit that the con-trol of chemotaxis by two (or more) pathways provides motilebacteria in a competitive environment. Given the ubiquitous dis-tribution of multiple chemotaxis pathways in the genomes ofrhizosphere bacteria, we expect this strategy to be widespread.

ACKNOWLEDGMENTS

This research is supported by National Science Foundation grant NSF-MCB 1330344 (to G.A.) and by the National Natural Science Foundationof China (31370108) and One Hundred-Talent Plan of the Chinese Acad-emy of Sciences (CAS) (to Z.X.).

Any opinions, findings, conclusions, or recommendations expressedin this material are those of the authors and do not necessarily reflect theviews of the National Science Foundation.

FUNDING INFORMATIONThis work, including the efforts of Zhi Hong Xie, was funded by NationalNatural Science Foundation of China (NSFC) (31370108). This work,including the efforts of Tanmoy Mukherjee, Dhivya Kumar, Nathan Bur-riss, Zhi Hong Xie, and Gladys Alexandre, was funded by National ScienceFoundation (NSF) (MCB-1330344).

REFERENCES1. Wadhams GH, Armitage JP. 2004. Making sense of it all: bacterial che-

motaxis. Nat Rev Mol Cell Biol 5:1024 –1037. http://dx.doi.org/10.1038/nrm1524.

2. Buchan A, Crombie B, Alexandre GM. 2010. Temporal dynamics andgenetic diversity of chemotactic-competent microbial populations in therhizosphere. Environ Microbiol 12:3171–3184. http://dx.doi.org/10.1111/j.1462-2920.2010.02290.x.

3. Wuichet K, Zhulin IB. 2010. Origins and diversification of a complex

signal transduction system in prokaryotes. Sci Signal 3:ra50. http://dx.doi.org/10.1126/scisignal.2000724.

4. Porter SL, Wadhams GH, Armitage JP. 2008. Rhodobacter sphaeroides:complexity in chemotactic signalling. Trends Microbiol 16:251–260. http://dx.doi.org/10.1016/j.tim.2008.02.006.

5. Zhulin IB, Armitage JP. 1993. Motility, chemokinesis, and methylation-independent chemotaxis in Azospirillum brasilense. J Bacteriol 175:952–958.

6. Bible A, Russell MH, Alexandre G. 2012. The Azospirillum brasilenseChe1 chemotaxis pathway controls swimming velocity, which affects tran-sient cell-to-cell clumping. J Bacteriol 194:3343–3355. http://dx.doi.org/10.1128/JB.00310-12.

7. Cecagno R, Fritsch TE, Schrank IS. 2015. The plant growth-promotingbacteria Azospirillum amazonense: genomic versatility and phytohormonepathway. Biomed Res 2015:7. http://dx.doi.org/10.1155/2015/898592.

8. Wisniewski-Dyé F, Lozano L, Acosta-Cruz E, Borland S, Drogue B,Prigent-Combaret C, Rouy Z, Barbe V, Herrera AM, González V,Mavingui P. 2012. Genome sequence of Azospirillum brasilense CBG497and comparative analyses of Azospirillum core and accessory genomesprovide insight into niche adaptation. Genes 3:576. http://dx.doi.org/10.3390/genes3040576.

9. Rivera D, Revale S, Molina R, Gualpa J, Puente M, Maroniche G, ParisG, Baker D, Clavijo B, McLay K, Spaepen S, Perticari A, Vazquez M,Wisniewski-Dyé F, Watkins C, Martínez-Abarca F, Vanderleyden J,Cassán F. 2014. Complete genome sequence of the model rhizospherestrain Azospirillum brasilense Az39, successfully applied in agriculture.Genome Announc 2(4):e00683-14. http://dx.doi.org/10.1128/genomeA.00683-14.

10. Wisniewski-Dyé F, Borziak K, Khalsa-Moyers G, Alexandre G,Sukharnikov LO, Wuichet K, Hurst GB, McDonald WH, Robertson JS,Barbe V, Calteau A, Rouy Z, Mangenot S, Prigent-Combaret C, Nor-mand P, Boyer M, Siguier P, Dessaux Y, Elmerich C, Condemine G,Krishnen G, Kennedy I, Paterson AH, González V, Mavingui P, ZhulinIB. 2011. Azospirillum genomes reveal transition of bacteria from aquaticto terrestrial environments. PLoS Genet 7:e1002430. http://dx.doi.org/10.1371/journal.pgen.1002430.

11. Stephens BB, Loar SN, Alexandre G. 2006. Role of CheB and CheR in thecomplex chemotactic and aerotactic pathway of Azospirillum brasilense. JBacteriol 188:4759 – 4768. http://dx.doi.org/10.1128/JB.00267-06.

12. Bible AN, Khalsa-Moyers GK, Mukherjee T, Green CS, Mishra P,Purcell A, Aksenova A, Hurst GB, Alexandre G. 2015. Metabolic adap-tations of Azospirillum brasilense to oxygen stress by cell-cell clumping andflocculation. Appl Environ Microbiol 81:8346 – 8357. http://dx.doi.org/10.1128/AEM.02782-15.

13. Jiang ZY, Gest H, Bauer CE. 1997. Chemosensory and photosensoryperception in purple photosynthetic bacteria utilize common signal trans-duction components. J Bacteriol 179:5720 –5727.

14. Hauwaerts D, Alexandre G, Das SK, Vanderleyden J, Zhulin IB. 2002.A major chemotaxis gene cluster in Azospirillum brasilense and relation-ships between chemotaxis operons in -proteobacteria. FEMS MicrobiolLett 208:61– 67. http://dx.doi.org/10.1111/j.1574-6968.2002.tb11061.x.

15. Vanstockem M, Michiels K, Vanderleyden J, Van Gool AP. 1987.Transposon mutagenesis of Azospirillum brasilense and Azospirillum li-poferum: physical analysis of Tn5 and Tn5-Mob insertion mutants. ApplEnviron Microbiol 53:410 – 415.

16. Higuchi R, Krummel B, Saiki RK. 1988. A general method of in vitropreparation and specific mutagenesis of DNA fragments: study of proteinand DNA interactions. Nucleic Acids Res 16:7351–7367. http://dx.doi.org/10.1093/nar/16.15.7351.

17. Krause A, Doerfel A, Gottfert M. 2002. Mutational and transcriptionalanalysis of the type III secretion system of Bradyrhizobium japonicum. MolPlant Microbe Interact 15:1228 –1235. http://dx.doi.org/10.1094/MPMI.2002.15.12.1228.

18. Alexandre G, Greer SE, Zhulin IB. 2000. Energy taxis is the dominantbehavior in Azospirillum brasilense. J Bacteriol 182:6042– 6048. http://dx.doi.org/10.1128/JB.182.21.6042-6048.2000.

19. Siuti P, Green C, Edwards AN, Doktycz MJ, Alexandre G. 2011. Thechemotaxis-like Che1 pathway has an indirect role in adhesive cell prop-erties of Azospirillum brasilense. FEMS Microbiol Lett 323:105–112. http://dx.doi.org/10.1111/j.1574-6968.2011.02366.x.

20. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ. 1990. Basic localalignment search tool. J Mol Biol 215:403– 410. http://dx.doi.org/10.1016/S0022-2836(05)80360-2.

Chemotaxis Signal Transduction in Azospirillum

June 2016 Volume 198 Number 12 jb.asm.org 1771Journal of Bacteriology

on Decem

ber 11, 2020 by guesthttp://jb.asm

.org/D

ownloaded from

21. Mohari B, Licata NA, Kysela DT, Merritt PM, Mukhopadhay S, BrunYV, Setayeshgar S, Fuqua C. 2015. Novel pseudotaxis mechanismsimprove migration of straight-swimming bacterial mutants through aporous environment. mBio 6:e00005. http://dx.doi.org/10.1128/mBio.00005-15.

22. Wolfe AJ, Berg HC. 1989. Migration of bacteria in semisolid agar. ProcNatl Acad Sci U S A 86:6973– 6977. http://dx.doi.org/10.1073/pnas.86.18.6973.

23. Miller LD, Yost CK, Hynes MF, Alexandre G. 2007. The major che-motaxis gene cluster of Rhizobium leguminosarum bv. viciae is essential forcompetitive nodulation. Mol Microbiol 63:348 –362. http://dx.doi.org/10.1111/j.1365-2958.2006.05515.x.

24. Bible AN, Stephens BB, Ortega DR, Xie Z, Alexandre G. 2008.Function of a chemotaxis-like signal transduction pathway in modu-lating motility, cell clumping, and cell length in the alphaproteobacte-rium Azospirillum brasilense. J Bacteriol 190:6365– 6375. http://dx.doi.org/10.1128/JB.00734-08.

25. Lauga E, DiLuzio WR, Whitesides GM, Stone HA. 2006. Swimming incircles: motion of bacteria near solid boundaries. Biophys J 90:400 – 412.http://dx.doi.org/10.1529/biophysj.105.069401.

26. Lemelle L, Palierne J-F, Chatre E, Place C. 2010. Counterclockwisecircular motion of bacteria swimming at the air-liquid interface. J Bacte-riol 192:6307– 6308. http://dx.doi.org/10.1128/JB.00397-10.

27. Armitage JP, Schmitt R. 1997. Bacterial chemotaxis: Rhodobacter spha-eroides and Sinorhizobium meliloti—variations on a theme? Microbiology143:3671–3682. http://dx.doi.org/10.1099/00221287-143-12-3671.

28. Greck M, Platzer J, Sourjik V, Schmitt R. 1995. Analysis of a chemotaxisoperon in Rhizobium meliloti. Mol Microbiol 15:989 –1000. http://dx.doi.org/10.1111/j.1365-2958.1995.tb02274.x.

29. Koonin EV, Makarova KS, Aravind L. 2001. Horizontal gene transfer inprokaryotes: quantification and classification. Annu Rev Microbiol 55:709 –742. http://dx.doi.org/10.1146/annurev.micro.55.1.709.

30. Porter SL, Wadhams GH, Armitage JP. 2011. Signal processing in com-plex chemotaxis pathways. Nat Rev Microbiol 9:153–165. http://dx.doi.org/10.1038/nrmicro2505.

31. Porter SL, Warren AV, Martin AC, Armitage JP. 2002. The thirdchemotaxis locus of Rhodobacter sphaeroides is essential for chemotaxis.Mol Microbiol 46:1081–1094. http://dx.doi.org/10.1046/j.1365-2958.2002.03218.x.

32. Shah DSH, Porter SL, Martin AC, Hamblin PA, Armitage JP. 2000. Finetuning bacterial chemotaxis: analysis of Rhodobacter sphaeroides behav-iour under aerobic and anaerobic conditions by mutation of the majorchemotaxis operons and cheY genes. EMBO J 19:4601– 4613. http://dx.doi.org/10.1093/emboj/19.17.4601.

33. Pilizota T, Brown MT, Leake MC, Branch RW, Berry RM, Armitage JP.

2009. A molecular brake, not a clutch, stops the Rhodobacter sphaeroidesflagellar motor. Proc Natl Acad Sci U S A 106:11582–11587. http://dx.doi.org/10.1073/pnas.0813164106.

34. Porter SL, Wadhams GH, Martin AC, Byles ED, Lancaster DE, Armit-age JP. 2006. The CheYs of Rhodobacter sphaeroides. J Biol Chem 281:32694 –32704. http://dx.doi.org/10.1074/jbc.M606016200.

35. Russell MH, Bible AN, Fang X, Gooding JR, Campagna SR, GomelskyM, Alexandre G. 2013. Integration of the second messenger c-di-GMPinto the chemotactic signaling pathway. mBio 4:e00001-13. http://dx.doi.org/10.1128/mBio.00001-13.

36. Stocker R, Seymour JR. 2012. Ecology and physics of bacterial che-motaxis in the ocean. Microbiol Mol Biol Rev 76:792– 812. http://dx.doi.org/10.1128/MMBR.00029-12.

37. Carvalhais LC, Dennis PG, Fan B, Fedoseyenko D, Kierul K, Becker A,von Wiren N, Borriss R. 2013. Linking plant nutritional status to plant-microbe interactions. PLoS One 8:e68555. http://dx.doi.org/10.1371/journal.pone.0068555.

38. Philippot L, Raaijmakers JM, Lemanceau P, van der Putten WH. 2013.Going back to the roots: the microbial ecology of the rhizosphere. Nat RevMicrobiol 11:789 –799. http://dx.doi.org/10.1038/nrmicro3109.

39. Hinsinger P, Bengough AG, Vetterlein D, Young I. 2009. Rhizosphere:biophysics, biogeochemistry and ecological relevance. Plant Soil 321:117–152. http://dx.doi.org/10.1007/s11104-008-9885-9.

40. Flessa H. 1994. Plant-induced changes in the redox potential of the rhi-zospheres of the submerged vascular macrophytes Myriophyllum verticil-latum L. and Ranunculus circinatus L. Aquat Bot 47:119 –129. http://dx.doi.org/10.1016/0304-3770(94)90009-4.

41. Greer-Phillips SE, Stephens BB, Alexandre G. 2004. An energy taxistransducer promotes root colonization by Azospirillum brasilense. J Bac-teriol 186:6595– 6604. http://dx.doi.org/10.1128/JB.186.19.6595-6604.2004.

42. Simon R, Priefer U, Puhler A. 1983. A broad host range mobilizationsystem for in vivo genetic engineering: transposon mutagenesis in gramnegative bacteria. Nat Biotechnol 1:784 –791. http://dx.doi.org/10.1038/nbt1183-784.

43. Keen NT, Tamaki S, Kobayashi D, Trollinger D. 1988. Improved broad-host-range plasmids for DNA cloning in Gram-negative bacteria. Gene70:191–197. http://dx.doi.org/10.1016/0378-1119(88)90117-5.

44. Dennis JJ, Zylstra GJ. 1998. Plasposons: modular self-cloning minitrans-poson derivatives for rapid genetic analysis of gram-negative bacterialgenomes. Appl Environ Microbiol 64:2710 –2715.

45. Töpfer R, Schell J, Steinbiss HH. 1988. Versatile cloning vectors fortransient gene expression and direct gene transfer in plant cells. NucleicAcids Res 16:8725. http://dx.doi.org/10.1093/nar/16.17.8725.

Mukherjee et al.

1772 jb.asm.org June 2016 Volume 198 Number 12Journal of Bacteriology

on Decem

ber 11, 2020 by guesthttp://jb.asm

.org/D

ownloaded from