JOURNAL OF BACTERIOLOGY, Nov. 2007, p. 8005–8014
Copyright © 2007, American Society for Microbiology. All Rights Reserved.
Vol. 189, No. 22
Motility and Chemotaxis in Agrobacterium tumefaciens Surface
Attachment and Biofilm Formation?†
Peter M. Merritt, Thomas Danhorn, and Clay Fuqua*
Department of Biology, Indiana University, Bloomington, Indiana 47405
Received 12 April 2007/Accepted 24 August 2007
Bacterial motility mechanisms, including swimming, swarming, and twitching, are known to have important
roles in biofilm formation, including colonization and the subsequent expansion into mature structured
surface communities. Directed motility requires chemotaxis functions that are conserved among many bacte-
rial species. The biofilm-forming plant pathogen Agrobacterium tumefaciens drives swimming motility by
utilizing a small group of flagella localized to a single pole or the subpolar region of the cell. There is no
evidence for twitching or swarming motility in A. tumefaciens. Site-specific deletion mutations that resulted in
either aflagellate, flagellated but nonmotile, or flagellated but nonchemotactic A. tumefaciens derivatives were
examined for biofilm formation under static and flowing conditions. Nonmotile mutants were significantly
deficient in biofilm formation under static conditions. Under flowing conditions, however, the aflagellate
mutant rapidly formed aberrantly dense, tall biofilms. In contrast, a nonmotile mutant with unpowered flagella
was clearly debilitated for biofilm formation relative to the wild type. A nontumbling chemotaxis mutant was
only weakly affected with regard to biofilm formation under nonflowing conditions but was notably compro-
mised in flow, generating less adherent biomass than the wild type, with a more dispersed cellular arrange-
ment. Extragenic suppressor mutants of the chemotaxis-impaired, straight-swimming phenotype were readily
isolated from motility agar plates. These mutants regained tumbling at a frequency similar to that of the wild
type. Despite this phenotype, biofilm formation by the suppressor mutants in static cultures was significantly
deficient. Under flowing conditions, a representative suppressor mutant manifested a phenotype similar to yet
distinct from that of its nonchemotactic parent.
The ability of bacteria to propel themselves through their
environment, often in response to chemical and physical gra-
dients, is an important adaptive mechanism that promotes
optimal positioning of cells at microscopic spatial scales. Gen-
erally, motile bacteria move through fluids and along surfaces
by using external appendages, including pili and flagella. Ro-
tation of single or multiple flagella drives swimming motility in
fluid environments and swarming along wetted surfaces (4).
Escherichia coli, with the best-understood model system of
bacterial motility, moves unidirectionally, with periodic pauses
that involve active tumbling, which reorient the cell prior to
continued forward motion (4). This essentially random move-
ment can be biased through taxis mechanisms that modify the
frequency of reorientation. Motility by other species, including
members of the Alphaproteobacteria, can differ significantly
from the E. coli paradigm, although other species often employ
conserved mechanisms (40). Type IV pili also mediate cellular
movement along surfaces, in a process called twitching motil-
ity, by directional extension and subsequent depolymerization
of the pili (25). Other microbes utilize gliding and ratchet-like
mechanisms to move on surfaces (27).
Motility can have a profound impact on the colonization of
surfaces, the first step in the formation of adherent microbial
assemblies called biofilms. Subsequent accumulation and lat-
eral expansion of adherent biomass can also involve motility. It
has been shown that aflagellate (Fla?) E. coli mutants do not
attach to surfaces in static culture (33). Active motility is re-
quired for adherence, as nonmotile E. coli mutants that make
flagella but cannot rotate them (Mot?) are also deficient in
biofilm formation (33). Mutations that disrupt flagellar biosyn-
thesis and twitching motility significantly alter biofilm forma-
tion and architecture in several pseudomonads (21, 32). Serra-
tia marcescens requires swarming motility via flagella and
surfactant production to effectively colonize surfaces (23). The
ability to rapidly spread across surfaces after initial attachment,
which is dependent on both flagella and pili, provides Pseudo-
monas aeruginosa with a significant competitive advantage in
mixed-species biofilms (2). Less direct evidence exists for the
impact of chemotaxis on surface interactions. Mutations that
disrupt the chemotaxis sensor kinase cheA, resulting in nontum-
bling mutants, do not affect biofilm formation by E. coli. Similar
chemotaxis mutants of Aeromonas spp. are reported to manifest
severe biofilm defects (20, 33). Colonization of surfaces as a
monolayer by Vibrio cholerae also involves chemotaxis (31). In
these systems, the mechanistic basis for the chemotaxis require-
ment in biofilm formation remains unclear.
Agrobacterium tumefaciens is a member of the Alphapro-
teobacteria that forms complex biofilms on abiotic surfaces and
plant tissues (10, 34). A. tumefaciens is best known as the
causative agent of crown gall, a neoplastic disease of plants.
Pathogenesis involves the horizontal transmission of a segment
of A. tumefaciens DNA, carried on the tumor-inducing (Ti)
plasmid, into the host plant genome (14). Genes on the trans-
ferred DNA direct a hormonal imbalance in the plant that
* Corresponding author. Mailing address: Department of Biology,
Indiana University, 1001 E. 3rd St., Jordan Hall 142, Bloomington, IN
47405-1847. Phone: (812) 856-6005. Fax: (812) 855-6705. E-mail:
† Supplemental material for this article may be found at http://jb
?Published ahead of print on 31 August 2007.
results in uncontrolled proliferation of plant cells, forming the
characteristic gall. Other transferred DNA genes direct the
synthesis of specialized nutrients by the plant, which are con-
sumed specifically by A. tumefaciens via catabolic pathways also
encoded on the Ti plasmid. Intense efforts have been focused
on the mechanics of this cross-kingdom genetic exchange, and
the process has been harnessed effectively in plant transgenesis
(8, 11, 14). However, our understanding of the activities that
lead to plant association and productive attachment is far more
limited. It is also important that many agrobacteria are present
in the bulk soil as avirulent plant-associated commensals (5).
Motility and chemotaxis have been implicated in plant associ-
ation and the early steps of disease (6, 37), but a thorough
analysis of their role in surface interactions has not been re-
Swimming motility in the most commonly studied strain of
A. tumefaciens, C58, is mediated by flagella, and there is no
evidence of swarming or twitching motility (17). Multiple fla-
gella are typically localized as a small tuft positioned at or
around a single pole of the cell (6). Similar to the well-studied
flagellar system of the related bacterium Sinorhizobium meliloti,
the flagella are thought to rotate in a clockwise direction to
propel the cell forward (3, 36). The current model states that
tumbling is not caused by a reversal of flagellar rotation, as in
the E. coli paradigm, but rather occurs through asynchronous
slowing of flagellar rotation, resulting in chaotic motion of the
flagella (3, 36). Flagellar assembly genes are carried in a single
major cluster of genes on the A. tumefaciens C58 circular
chromosome (Atu0542 to Atu0582), including four discrete
flagellin gene homologues (15, 45). Mutants deleted for three
of the four flagellin genes (?flaABC) are reported to be non-
motile and are attenuated for virulence (6). The genes for
several motor (Mot) proteins, homologues of which drive
flagellar rotation in other systems, are also located within this
cluster, although these genes and their role in motility have not
been studied thus far.
The A. tumefaciens C58 genome also encodes roughly 20
methyl-accepting chemotaxis protein (MCP) homologues, sug-
gesting complex adaptive control over motility (15, 45). The
main chemotaxis functions, including a single cheA gene en-
coding a two-component sensor kinase that interacts with the
MCPs and two discrete response regulator genes (cheY1 and
cheY2), are located within the Che operon (Atu0514 to
Atu0522), which is near the flagellar assembly cluster (46).
This operon is syntenous to that found in S. meliloti, with the
exception that cheW is not present (13). In contrast, there are
two cheW homologues at separate positions on the C58 circular
chromosome (Atu2075 and Atu2617). As in S. meliloti, there is
no cheZ phosphatase gene homologue present in strain C58. A
nonpolar cheA deletion mutant exhibited a smaller swim ring
in motility agar and was less efficient than the wild type for
tumorigenesis (46). Chemotaxis by A. tumefaciens towards phe-
nolic plant exudates and opines has been demonstrated, and
mutation of an MCP-type protein gene on the Ti plasmid
abolished chemotaxis towards opines (19, 37).
In this study, we have generated and analyzed defined A.
tumefaciens C58 mutants that are aflagellate (?flgE), flagel-
lated but nonmotile (?motA), and motile but nonchemotactic
(?cheA) for biofilm formation on abiotic surfaces in static
culture and in flow cells. In so doing, we have also isolated an
intriguing spontaneous suppressor of the chemotaxis mutation
that reestablishes the ability to tumble but severely compro-
mises biofilm formation. Our findings demonstrate important
roles for chemotaxis and motility in biofilm formation under
flowing and static conditions and show that biofilm attributes
impacted by these functions can vary depending on prevailing
MATERIALS AND METHODS
Strains, plasmids, and growth conditions. All bacterial strains and plasmids
used in this study are listed in Table 1. Reagents, antibiotics, and microbiological
media were obtained from Fisher Scientific (Pittsburgh, PA) and Sigma-Aldrich
(St. Louis, MO). DNA manipulations were performed following standard pro-
tocols (35), and DNA sequences were determined on an ABI 3730 sequencer
(Indiana Molecular Biology Institute, Bloomington, IN). Oligonucleotide prim-
ers were supplied by Integrated DNA Technologies, Coralville, IA (primer
information is listed in Table 2). Nucleic acid purification was performed using
QIAquick Spin kits (QIAGEN, Valencia, CA). Plasmids were introduced into A.
tumefaciens by electroporation (28) or conjugal transfer (12). Bacteria were
maintained on either LB or AT minimal medium (39) supplemented with 0.5%
(wt/vol) glucose and 15 mM ammonium sulfate (ATGN). To prevent the accu-
mulation of iron oxide precipitate, the FeSO4? 7H2O prescribed in the original
AT recipe was omitted, with no adverse growth effect. For sacB counterselection,
0.5% sucrose (Suc) replaced glucose as the sole carbon source (ATSN). Antibi-
otic concentrations used were as follows: for E. coli, 100 ?g ml?1ampicillin, 25
?g ml?1streptomycin (Sm), and 25 ?g ml?1kanamycin (Km); and for A.
tumefaciens, 3 mg ml?1Sm, 150 ?g ml?1Km, and 50 ?g ml?1spectinomycin, as
Construction of nonpolar markerless deletions. To generate nonpolar dele-
tions, PCR was used to amplify approximately 500 bp of flanking sequence
upstream (primers 1 and 2) and downstream (primers 3 and 4) of the reading
frame to be deleted. Primers were designed to remove as much of the coding
sequence as possible without disrupting any predicted translational coupling.
TABLE 1. Strains and plasmids used in this study
Strain or plasmidRelevant feature(s)
C58 Nopaline type strain; pTiC58,
?cheA C58 derivative
?flgE C58 derivative
?motA C58 derivative
Spontaneous suppressor of
E. coli strains
?pir Tra?; cloning strain
?pir; cloning strain
PCR cloning vector; Apr
R6K ori; sacB (Sucs) Smr
Ptac::gfpmut3 Spr; pVS replicon
pKNG101 carrying cheA SOE
pKNG101 carrying flgE SOE
pKNG101 carrying motA SOE
J. Zhu (9)
8006 MERRITT ET AL.J. BACTERIOL.
Primers 2 and 3 were designed with 18-bp complementary sequences at their 5?
ends (lowercase nucleotides in Table 2) to facilitate splicing by overlapping
extension (SOE), essentially as described previously (41). Briefly, both flanking
sequences were amplified using the high-fidelity Phusion DNA polymerase
(NEB, Beverly, MA) and were agarose gel purified. Purified PCR products were
used as both templates and primers for a five-cycle PCR. A final PCR step with
primers 1 and 4, using 2 ?l of the second-step reaction mix as the template,
generated the full-length spliced product. The final PCR products were cloned
into pGEM-T Easy (Promega, Madison, WI), confirmed by sequencing, excised
by cleavage with the appropriate restriction enzyme, and ligated with the suicide
vector pKNG101 cleaved at compatible restriction sites. The pKNG101 plasmid
confers Sm resistance (Smr) and sucrose sensitivity (Sucs) (18). Derivatives of
pKNG101 were introduced into A. tumefaciens C58 by conjugal transfer. Because
the R6K origin of pKNG101 does not replicate in A. tumefaciens, the plasmid
must recombine into the genome to allow growth in media containing Sm.
Recombinants were selected on ATGN plates containing Sm, and plasmid inte-
gration was confirmed by patching Smrisolates onto ATSN-Sm plates to identify
Sucsclones. To facilitate excision of the integrated plasmid, SmrSucsclones were
grown overnight in ATGN broth without Sm and then plated on ATSN. Plasmid
excision was verified by patching Sucrclones onto ATSN plus Sm to identify Sms
isolates. Appropriate deletion of the target genes was confirmed by diagnostic
PCR and DNA sequencing of the products (with primers 5 and 6, which flank the
Complementation constructs. To perform complementation analyses, wild-
type coding sequences were cloned into pBBR1MCS-2 (22). 5? primers were
designed with a stop codon in frame with pBBR-carried lacZ followed by the E.
coli lacZ ribosome binding site to prevent translational occlusion and to optimize
expression, respectively (Table 2). Coding sequences for cheA (Atu0517), flgE
(Atu0574), and motA (Atu0560) were PCR amplified from C58 genomic DNA,
using the corresponding primers 7 and 8 for each gene (Table 2) and the Phusion
polymerase, ligated into pGEM-T Easy, confirmed by sequencing, excised by restric-
tion enzyme cleavage, and ligated with appropriately cleaved pBBR1MCS-2. Plas-
mid derivatives harboring the correct inserts were verified by restriction digestion
and sequencing prior to electroporation into competent A. tumefaciens cells.
Flagellar staining and motility assays. A flagellar staining protocol was
adapted from the work of Mayfield and Inniss (26). Briefly, a two-component
stain [solution A is equal volumes 5% phenol and saturated AlK(SO4)2? 12H2O
in 10% tannic acid, and solution B is 12% crystal violet (CV) in 100% ethanol]
was mixed fresh at a ratio of 10 volumes solution A to 1 volume solution B,
vortexed, and centrifuged to remove CV crystals. One microliter of cell culture
was spotted onto a clean microscope slide and overlaid with a 22- by 22-mm
coverslip. The slides were held vertically, and approximately 2 to 5 ?l of stain was
applied to the edge of the coverslip. Capillary action wicked the stain under the
coverslip, staining the flagella. Samples were observed by phase-contrast micros-
copy using a 100? oil immersion objective after at least 5 min of staining.
Swimming and chemotaxis phenotypes were tested on ATGN swim agar plates
containing 0.25% Bacto agar (BD, Sparks, MD) (1). Two plating formats were
used. First, 100-mm petri plates were filled with 25 ml ATGN swim agar. Swim
plates were inoculated from fresh colonies or cultures by using a toothpick that
was stabbed into the agar at the center of the plate. Second, for screening
purposes, 12-well tissue culture plates (Corning, Corning, NY) were filled with 3
ml swim agar per well and inoculated as described above.
Short-term binding assays. Short-term binding assays were conducted by
growing the appropriate green fluorescent protein (GFP)-expressing strains in
ATGN to an optical density at 600 nm (OD600) of approximately 0.6. Glass
coverslips were floated on 5 ml of culture in six-well tissue culture plates for 2 to
4 hours. Coverslips were removed from the plates, rinsed thoroughly with 1? AT
buffer (79 mM KH2PO4, pH 7.0), and mounted on slides for microscopy. Each
strain was tested in triplicate, with 10 fields of view captured for each coverslip
by spinning disk confocal microscopy (Yokagawa CSU10 confocal scanner unit,
Nikon TE2000U microscope, and Photometrics Cascade II 512B camera) with a
40? objective, using MetaMorph software (Molecular Devices Corp., Sunnyvale,
CA). Cells were counted manually or automatically using ImageJ (NIH).
Cultivation and analysis of static culture biofilms. Static culture biofilms were
grown essentially as described previously (34). Briefly, polyvinyl chloride (PVC)
coverslips were placed vertically in 12-well polystyrene cell culture plates (Corn-
ing Inc.), inoculated with cells in ATGN at an OD600of 0.05, and incubated at
room temperature for 24 to 96 h. Biofilms were visualized macroscopically by CV
staining or microscopically by phase-contrast and epifluorescence microscopy.
For CV visualization, coverslips were rinsed in double-distilled H2O, stained with
0.1% (wt/vol) CV, and rinsed again in double-distilled H2O. Biomass adhered to
the coverslip was quantified by soaking stained coverslips in 1 ml dimethyl
sulfoxide (DMSO) to solubilize the CV and measuring the absorbance at 600 nm
(A600) in a Bio-Tek Synergy HT microplate reader. Absorbance values were
normalized to culture growth by dividing the A600value for solubilized CV by the
OD600of the planktonic cells.
Cultivation and analysis of flow cell biofilms. Once-through flow cells with a
200-?l chamber volume were inoculated with A. tumefaciens carrying pJZ383
(Ptac::gfpmut3) for GFP expression, as described previously (7, 10). All tubing
and bubble traps were autoclaved prior to assembly of the flow cell system. The
system was filled with 0.5% sodium hypochlorite and left overnight without flow.
A minimum of 2 liters of sterile water was flushed through the system prior to
treatment with 0.6% hydrogen peroxide at a flow rate of ?30 ml h?1for at least
3 h. Flow cells were flushed with 2 to 3 liters of sterile water and then equilibrated
with ATGN (flow rate of 3 ml h?1) for at least 12 h prior to inoculation. For each
strain tested, three individual flow channels were inoculated. Each chamber was
inoculated with 200 ?l of cells suspended in ATGN at an OD600of 0.05. After 1
hour, flow was resumed at a rate of 3 ml h?1and continued uninterrupted for the
duration of the experiment. Five image stacks per channel were acquired on a
spinning disk confocal microscope using MetaMorph software. Each field of view
had an area of approximately 1.45 ? 104?m2. Stacks were processed and
analyzed using autoCOMSTAT, a modified version of the COMSTAT biofilm evalua-
tion package by Heydorn et al. (16; T. Danhorn and C. Fuqua, unpublished
data). Biofilm parameters measured in this study were biovolume (?m3per ?m2
substatum area), substratum coverage (% within the first micrometer above the
substratum), average height (?m), roughness (a unitless measure of height vari-
ability), and the total number of microcolonies counted in 15 fields of view.
Deletion mutations in motility and chemotaxis genes. To
investigate the role of flagellar motility in biofilm formation,
nonpolar, markerless deletions were generated in key genes en-
coding flagellar biosynthesis (Fla?), flagellar rotation (Mot?),
TABLE 2. Primer sequences
aEngineered restriction sites are shown in lowercase. Underlining highlights
the features engineered for optimized expression from the pBBR1-MCS2 vector
(see Materials and Methods).
VOL. 189, 2007 BIOFILMS AND BACTERIAL LOCOMOTION8007
and chemotaxis (Che?) proteins. Precise deletions were gen-
erated in the A. tumefaciens C58 genes encoding the flagellar
hook protein (flgE; Atu0574), a component of the flagellar
motor (motA; Atu0560), and the chemotaxis sensor kinase
(cheA; Atu0517). No motile ?flgE or ?motA cells were ob-
served microscopically, while the ?cheA mutant was motile but
nontumbling (data not shown). Flagellar staining of the non-
motile mutants showed, as expected, that the ?flgE mutant was
aflagellate, while the ?motA strain was flagellated (Fig. 1).
Flagellar assembly and swimming were restored when wild-
type copies of the deleted genes were expressed in trans (Fig.
1E and data not shown). All mutants grew at rates similar to
that of the wild type in aerated cultures (data not shown).
Motility mutant swimming phenotypes were assessed on
0.25% swim agar plates. Neither the ?flgE nor the ?motA
mutant expanded beyond the site of inoculation (Fig. 2B and
C). Consistent with other A. tumefaciens Che?mutants (46),
our ?cheA mutant formed dense, small-diameter swim rings, in
contrast to the diffuse, large-diameter swim rings observed in
the wild type (Fig. 2D). For all of the mutants, normal swim
phenotypes were restored by provision of a plasmid-borne copy
of the deleted gene (Fig. 2E, F, and G).
Flagellar mutants are significantly compromised for attach-
ment and biofilm formation. The C58 motility mutants were
monitored for attachment and biofilm formation on PVC cov-
erslips in ATGN. Under these conditions, the ?flgE and
?motA mutants were significantly deficient in their ability to
form biofilms. The CV extracted from stained biofilms showed
a ?50% reduction of attached biomass for both mutants (Fig.
3A). Nonmotile mutants exhibited growth rates similar to that
of the wild type in static cultures (data not shown). We con-
ducted short-term binding assays to distinguish impaired at-
tachment from deficiencies in subsequent biofilm maturation.
In these assays, glass coverslips were floated on a dense sus-
pension of cells for several hours and imaged by spinning disk
confocal microscopy, and attached cells were counted. In these
assays, the nonmotile mutants manifested a 90% reduction in
attachment (Fig. 3B).
CV-stained ?flgE biofilms on PVC coverslips manifested a
macroscopic punctate pattern distinct from the pattern for
wild-type biofilms (data not shown). Examination of coverslip
biofilms grown with GFP-expressing ?flgE mutants by epiflu-
FIG. 1. Flagellar staining phenotypes of motility mutants. Phase-
contrast images of CV-stained flagella from wild-type C58 and motility
mutants are shown. Images were taken with a 100? objective. (A) C58
wild type; (B) ?flgE mutant; (C) ?motA mutant; (D) ?cheA mutant;
(E) ?flgE/pPM110 mutant; (F) cms-1 mutant.
FIG. 2. Swim plate assays of C58 motility mutant and comple-
mented mutant strains. ATGN plates supplemented with 0.25% agar
were inoculated with wild-type C58 (A), the ?flgE mutant (B), the
?motA mutant (C), the ?cheA mutant (D), the ?flgE/pPM110 mutant
(E), the ?motA/pPM111 mutant (F), and the ?cheA/pPM109 mutant (G).
FIG. 3. Static biofilm formation and surface attachment by C58
motility mutants. (A) Quantification of DMSO-solubilized CV from
PVC coverslip biofilms at 72 hpi. Biomass normalized for growth
(A600/OD600; closed bars) and adherent biomass (CV A600; open bars)
are shown. ?, strains carrying a wild-type copy of the deleted gene on
pBBR1MCS-2 (Table 1). Error bars show standard errors of the means
(SEM) for three coverslips. (B) Static culture short-term binding assay.
Total numbers of attached cells per field of view (?3.13 ? 104?m2) are
shown. Values are averages of 30 fields of view. Error bars show SEM.
8008MERRITT ET AL. J. BACTERIOL.
orescence microscopy revealed fewer attached cells but greater
numbers of microcolonies than those in wild-type C58 biofilms
(data not shown). The ?motA mutant tended to manifest a
more severe biofilm defect than the ?flgE mutant, although
this was not significant in all experiments (data not shown).
Plasmid-borne copies of the deleted wild-type genes comple-
mented each mutant biofilm phenotype.
Chemotaxis influences A. tumefaciens C58 attachment and
biofilm formation in static culture. The ?cheA mutant re-
vealed a modest reduction in static culture biofilm formation
relative to the wild type (Fig. 3A). However, when CV staining
was normalized for culture growth (CV absorbance divided by
planktonic culture turbidity), the relative amounts of adherent
biomass were similar (Fig. 3A, gray bars). Static culture growth
curves performed in the same format as the biofilm assays
revealed a significant growth defect in the ?cheA mutant that
confounded interpretation of the ?cheA biofilm phenotype
(data not shown). Normal growth was restored by a plasmid-
borne copy of wild-type cheA.
In order to assess surface interactions of the ?cheA mutant
independent of the growth rate, we conducted short-term bind-
ing assays as described above. At the time scale used in this
experiment, the effects of growth were minimal. In this assay,
the number of attached ?cheA cells was approximately 40% of
the wild-type cell number (Fig. 3B).
Robust biofilm formation in flow cells by ?flgE mutants. We
compared biofilm formation of the C58 motility mutants in
once-through flow chambers under low-shear conditions. Flow
cells are a continuous culture format in which the biofilm is
constantly irrigated with fresh medium. In contrast to the
marked biofilm deficiency observed in static assays, the ?flgE
mutant formed very thick and dense biofilms with numerous
large towers by 96 h postinoculation (hpi) (Fig. 4B; see Fig.
S1B in the supplemental material). After this point, the basic
morphology of ?flgE biofilms did not change. At 144 hpi,
wild-type C58 biofilms were qualitatively similar to those of the
?flgE mutant, indicating a significant acceleration in the rate of
biofilm formation for the mutant but no gross alterations in
biofilm architecture, aside from a pronounced difference in
height (see Fig. S1A and B in the supplemental material).
We used a modified version of the COMSTAT biofilm analysis
program (16), called autoCOMSTAT (Danhorn and Fuqua, un-
published data), to quantitatively analyze flow cell biofilms.
These analyses confirmed that the rate of ?flgE biofilm forma-
tion was greatly accelerated relative to that of the wild type. By
96 hpi, biofilms formed by the ?flgE mutant showed 5- to
15-fold increases in the total biovolume, percentage of substra-
tum coverage, and average height relative to those of wild-type
C58 (Fig. 5). The average maximum height from 15 fields of
view was approximately 50 ?m for ?flgE biofilms at 120 hpi, in
contrast to approximately 30 ?m for the wild type at 144 hpi.
The rate at which ?flgE biofilms accumulated increased rapidly
after 48 hpi, while a similar rate increase occurred only after 96
hpi in wild-type biofilms (Fig. 5). The maximum rates of bio-
film formation were similar between the wild type and the
?flgE mutant (compare ?flgE mutant rates at 48 to 96 hpi to
those of C58 at 96 to 144 hpi in Fig. 5). The ?flgE mutant
biofilms had saturated the flow cell system by 96 hpi, while
FIG. 4. Flow cell biofilms. Confocal laser scanning microscopy images of strain C58 and motility mutant derivatives expressing GFP grown in
ATGN are shown. (A) C58 at 144 hpi; (B) ?flgE mutant at 96 hpi; (C) ?motA mutant at 144 hpi; (D) ?cheA mutant at 144 hpi; (E) cms-1 mutant
at 144 hpi. Side and bottom panels are orthogonal views of the biofilms. Bar ? 20 ?m in all dimensions.
FIG. 5. Image analysis of flow cell biofilms. autoCOMSTAT was used to determine the biovolume per substratum area (A), substratum
coverage (B), and average overall height (C). Values are averages calculated for 15 image stacks collected from three channels (5 image stacks per
channel) for each strain per time point. Error bars show SEM.
VOL. 189, 2007BIOFILMS AND BACTERIAL LOCOMOTION 8009
wild-type biofilms continued to accumulate for the duration of
We also enumerated microcolonies, which we defined as
adherent cell aggregates with a volume of at least 100 ?m3.
The maximum number of microcolonies counted in the ?flgE
biofilms occurred at 48 hpi, corresponding to 20% substratum
coverage (Fig. 5B; see Fig. S2 in the supplemental material).
From 48 to 72 hpi, the number of microcolonies dropped
fourfold, reflecting the increasing confluence of adherent
growth. Wild-type biofilms reached a maximum number of
microcolonies at 96 hpi, 2 days later than the ?flgE mutant, but
with similar amounts of colonized substratum (Fig. 5B; see Fig.
S2 in the supplemental material). As the substratum coverage
approached 60% at 144 hpi, the number of C58 microcolonies
was approximately one per field of view, indicating nearly con-
tiguous coverage of the substratum.
To further investigate the nature of the ?flgE biofilm phe-
notype, we conducted a short-term flow cell experiment, mak-
ing observations every 3 hours. For the first 9 hours, the ?flgE
mutant biovolume and substratum coverage values were
slightly lower than those for the wild type, consistent with its
attachment deficiency in static cultures (see Fig. S3 in the
supplemental material). However, by 18 hpi, the ?flgE mutant
exceeded the wild type in both of these parameters. By 24 hpi,
the amounts of ?flgE biovolume and substratum coverage were
approximately 1.5- to 2-fold higher than those of wild-type C58
(see Fig. S3 in the supplemental material).
The ?motA mutant is severely compromised for biofilm for-
mation in flow cells. Similar to the profound ?motA mutant
biofilm defect in static culture, the ?motA mutant flow cell
biofilms were dramatically reduced relative to those of the wild
type. Early time points revealed limited numbers of attached
cells and isolated microcolonies. Although large microcolonies
had formed by 144 hpi, extensive areas of uncolonized substra-
tum remained (Fig. 4C; see Fig. S1C in the supplemental
material). The ?motA mutant biofilms were debilitated in sev-
eral of the numerical parameters analyzed, with approximately
threefold reductions in biovolume, substratum coverage, and
average height relative to those of the wild type by 120 hpi (Fig.
5). Even with these differences, the total biovolume and aver-
age height approached wild-type levels by 144 hpi (Fig. 5), and
both strains had similar maximum heights of approximately 30
?m. However, biofilms formed by the ?motA mutant had
about 30% less substratum coverage than the wild type (Fig. 5).
Consistent with this observation, the number of ?motA micro-
colonies continued to increase through 144 hpi, revealing that
the increases in total adherent biomass over time were con-
strained to specific sites on the surface (see Fig. S2 in the
Comparison of the average biofilm height and the roughness
coefficient is a metric that has been proposed to represent
biofilm complexity (16). The dimensionless roughness coeffi-
cient is a parameter that provides a measure of biofilm height
variability. By this comparison, biofilms formed by the ?motA
mutant maintained an architecture significantly different from
that of the wild type, with greater heterogeneity over the du-
ration of the experiment (Fig. 6C).
Chemotaxis is required for normal biofilm formation in flow
cells. The ?cheA mutant manifested an aberrant biofilm phe-
notype in flow cells that was more pronounced than that in
static culture biofilms. At early time points, very few cells were
attached to the substratum. However by 144 hpi, the biofilms
covered a substantial fraction of the available surface but were
relatively homogeneous and short (Fig. 4D; see Fig. S1D in the
supplemental material). The ?cheA mutant biofilms also ap-
peared to be less dense than those of the wild type or the
nonmotile mutants (Fig. 4B and C; see Fig. S1D in the sup-
plemental material). Qualitative observations were confirmed
by autoCOMSTAT analysis. The biomass and height of the ?cheA
mutant biofilms were roughly threefold lower than those of
C58 biofilms by 144 hpi, with less overall substratum coverage
(Fig. 5). Similar to the ?motA biofilms, the number of micro-
colonies in biofilms of the ?cheA mutant increased through
144 hpi (see Fig. S2 in the supplemental material). The com-
parison of average height to roughness reflected a distinct
architecture for the ?cheA biofilms throughout the experiment
Identification of a bypass suppressor of the ?cheA motility
agar phenotype. The motility agar phenotype of the ?cheA
mutant consistently exhibited an about 70% reduction in swim
ring diameter relative to that of wild-type C58. Interestingly,
after extended incubation, flares of diffuse growth began to
emerge from the dense swim ring formed by the ?cheA mutant
(Fig. 7A). With time, these flares expanded to nearly cover the
plate at a density similar to that of the wild type (data not
shown). Bacteria isolated from the outer edges of these flares
FIG. 6. Average height and roughness of flow cell biofilms at 48 (A), 96 (B), and 144 (C) hpi. Data are shown for C58 (closed squares), the
?cheA mutant (open squares), the ?motA mutant (open circles), and the cms-1 mutant (closed circles). Note that the ?flgE mutant was omitted
from these comparisons because of the large differences in biofilm height. The coordinates (height, roughness) for the ?flgE mutant were as
follows: 0.563 ?m, 1.532 at 48 hpi, 11.259 ?m, 0.762 at 96 hpi, and 11.137 ?m, 0.685 at 144 hpi. Error bars show SEM.
8010 MERRITT ET AL.J. BACTERIOL.
exhibited a swim phenotype similar to that of the wild type
(Fig. 7B). Quantitative analysis showed that swim ring diame-
ters of a representative suppressor mutant, the cms-1 mutant,
were approximately 80% that of strain C58 (Fig. 7D). All
putative suppressor isolates tested retained the cheA deletion.
Additional chemotaxis mutants, including a mutant in which
the entire chemotaxis operon was deleted (Atu0514 to
Atu0522), also generated similar spontaneous suppressor mu-
tations (data not shown). Microscopic examination revealed
appreciable tumbling in the suppressor mutants, in contrast to
the ?cheA nontumbling phenotype. We designated this the
Cms phenotype, for Che?mutation suppressor. When a copy
of cheA was expressed from a plasmid in the cms-1 mutant,
swim ring diameters were restored to normal (Fig. 7C and D).
cms mutants are impaired in biofilm formation in static
culture and flow cells. The cms mutants appeared to be nearly
wild type in swimming behavior. However, in contrast to the
wild type and their ?cheA parent strain, the cms mutants ex-
hibited profound biofilm formation deficiencies. In static cul-
ture coverslip biofilms, there was roughly 50% less attached
cms-1 biomass after 72 h of growth (Fig. 8A). Despite correct-
ing the Cms swim plate phenotype to full wild-type levels (Fig.
7C and D), plasmid-based expression of wild-type cheA did not
complement the biofilm defect (Fig. 8A). Similar to the case
for the other motility and chemotaxis mutants, the cms-1 mu-
tant manifested a severe attachment deficiency in short-term
binding assays (compare Fig. 8B to Fig. 3B).
Flow cell biofilms of the cms-1 mutant were dramatically
altered relative to those of the wild type (Fig. 4; see Fig. S1E
in the supplemental material). Although similar to the ?cheA
mutant biofilms, several of the attributes of the cms-1 biofilm
were also distinct. cms-1 biofilms attained an average height
similar to that of the wild type by 144 hpi but were more
structurally homogeneous and manifested significantly less sur-
face coverage and overall biomass (Fig. 4E and 5). Late-stage
cms-1 biofilms were also less dense, with a roughness coeffi-
cient approximately twofold higher than that of the wild type,
while both strains had similar average heights (Fig. 6C; also
compare the orthogonal views in Fig. 4A and E).
In this study, we examined biofilm formation by defined A.
tumefaciens C58 motility and chemotaxis mutants in both static
and flow cell cultivation formats. We observed that the roles of
motility and chemotaxis in biofilm formation differ significantly
between these culture formats. The defined Fla?and Mot?
mutants described here manifested severe deficiencies in at-
FIG. 7. Motility agar phenotype of the cms-1 mutant. Swim plate
images of the ?cheA mutant showing suppressor flare (A), the cms-1
mutant (B), and the cms-1/pPM109 mutant (C) are displayed. Image A
was taken at 6 days postinoculation; images B and C were taken at 5
days postinoculation. The arrow shows the suppressor flare from the
?cheA swim ring. (D) Quantified swim ring diameters of C58 (closed
squares), the ?cheA mutant (open squares), the ?flgE mutant (closed
triangles), the ?motA mutant (open triangles), the cms-1 mutant
(closed circles), and the cms-1/pPM109 mutant (open circles). Values
are the averages for four swim plates per strain. Error bars show SEM.
FIG. 8. Static biofilm formation and surface attachment by the
cms-1 mutant. (A) Quantification of DMSO-solubilized CV from PVC
coverslip biofilms at 72 hpi. Biomass normalized for growth (A600/
OD600; closed bars) and adherent biomass (CV A600; open bars) are
shown. ?, strains carrying a wild-type copy of the deleted gene on
pBBR1MCS-2. Error bars are SEM for three coverslips. (B) Static
culture short-term binding assay. Total numbers of attached cells per
field of view (?3.13 ? 104?m2) are shown. Values are averages for 30
fields of view. Error bars show SEM.
VOL. 189, 2007 BIOFILMS AND BACTERIAL LOCOMOTION 8011
tachment and biofilm formation in static culture. Interestingly,
the ?flgE mutant biofilms grown in flow cells were greatly
enhanced relative to those of the wild type, while the ?motA
biofilms remained severely debilitated. Chemotaxis mutants
were more modestly affected in attachment and biofilm forma-
tion under static conditions but manifested a more severe
defect in flow cells. Finally, a spontaneous suppressor mutation
that restores tumbling to the straight-swimming ?cheA mutant
had a negative impact on biofilm formation in both static and
flow cell cultures. The flow cell format allowed us to perform
high-resolution comparisons of the wild type and mutant de-
rivatives during the progression from initial attachment
through mature biofilms (see Fig. S1 in the supplemental ma-
Motility-driven surface interactions differentially influence
biofilm formation between static and flowing environments.
There have been few direct comparisons of biofilm formation
by motile bacteria between static and flowing formats, partic-
ularly those that evaluate the roles for chemotaxis and motility.
Under static culture conditions, a loss of motility should limit
the ability of cells to interact with and subsequently attach to
surfaces. Studies of E. coli Fla?and Mot?mutants suggested
that motility per se is required for biofilm formation and that
flagella do not function as inert adhesins (33). In contrast,
Kirov et al. reported that the flagella of Aeromonas spp. func-
tion primarily as adhesins that promote attachment (20). If A.
tumefaciens C58 flagella function strictly as adhesins, one
would expect the phenotype of ?motA mutation to be less
pronounced than that of ?flgE mutation. Our data clearly show
that depending on the culture format, the ?motA biofilm de-
fect is equal to or more severe than that of the ?flgE mutant,
suggesting that active motility is required for attachment and
biofilm maturation. Thus, flagella do not function as inert
adhesins in C58. A modest yet consistent trend in our static
biofilm assays suggests that the ?motA mutation imparts a
more severe defect than the ?flgE mutation (data not shown).
We therefore propose that the presence of unpowered flagella
in the ?motA mutant does in fact interfere with initial surface
interactions and subsequent permanent attachment.
We expected biofilm phenotypes of the motility mutants to
be less severe in flow cells because increased surface sampling,
imparted by the flow of medium, would allow for more fre-
quent attachment and subsequent biofilm formation. We were
surprised, however, to find that the accumulation of adherent
biomass was significantly accelerated for the C58 ?flgE mutant.
The adherent population was strikingly thicker than that of the
wild type and had significant numbers of very tall and dense
tower-like structures. Therefore, increasing the frequency of
surface interactions relative to that under static conditions
facilitates robust adherence and biofilm formation by the ?flgE
In contrast, flow conditions did not ameliorate the ?motA
attachment defect, suggesting that paralyzed flagella may phys-
ically interfere with surface attachment. A recent report has
shown that nonmotile Listeria monocytogenes mutants exhibit
profound deficiencies in surface attachment (24). For aflagel-
late mutants, this attachment defect can be rescued by forcing
cells to the surface via centrifugation, but this phenotype of
mutants with paralyzed flagella is only partially rescued. These
observations suggest that paralyzed flagella may block surface
interactions and thereby limit attachment efficiency and bio-
film formation. This is consistent with our findings that clearly
indicate that for strain C58, unpowered flagella are not effi-
cient adhesins in static or flowing environments and block
productive surface interactions. Another possibility is that the
flagella provide enhanced drag for the flowing medium to pull
the cells away from the surface.
Lateral biofilm expansion without surface motility mecha-
nisms. Biofilm expansion along surfaces involves a combina-
tion of multiple processes, including colonization from plank-
tonic-phase cells, clonal growth of surface-associated cells, and
in some systems, migration of previously attached cells via
surface motility. In P. aeruginosa, twitching and swarming mo-
tilities promote lateral expansion and contribute to the overall
architecture of the biofilm (21). Twitching and swarming have
not been reported for C58 or other agrobacteria. The lateral
accumulation of adherent biomass for C58 must therefore
combine clonal growth of attached cells and new colonization
by planktonic bacteria, the majority of which originate from
the biofilm itself.
For nonflagellated C58 mutants in static cultures, we ex-
pected that the small fraction of cells colonizing the surface
from the fluid phase would result in clonal microcolonies due
to reduced lateral spreading, leading to a patchy biofilm. The
punctate appearance of the ?flgE coverslip biofilms is consis-
tent with this prediction (data not shown). Flow cell biofilms of
this mutant were comprised of strikingly tall microcolonies and
rapidly formed dense surface populations (Fig. 4B and 5; see
Fig. S3 in the supplemental material). This aberrant micro-
colony formation seems readily explained by clonal growth of
adhered cells combined with limited dispersal. The overall
adherent biomass for the ?flgE mutant was, however, much
greater than that of the wild type (Fig. 5A), suggesting addi-
tional roles for motility subsequent to initial colonization and
For many motile bacteria, there is a period of reversible
attachment during which cells can detach and sample addi-
tional surfaces (43). It is likely that in the absence of swimming
motility, ?flgE mutant cells at the surface frequently transition
to irreversible attachment because there is no mechanism by
which the cells can actively detach. The frequency at which
bacteria detach from surfaces will have a significant impact on
the growth and architecture of the biofilm. In the wild type, the
rate of biomass increase on the surface is contingent upon
active emigration of motile cells into the planktonic phase. In
contrast, ?flgE cells cannot actively detach, thus constraining
their escape from the surface relative to that of the wild type.
This effect would grow to be more pronounced as the attached
biomass accumulates, thereby increasing the total number of
cells near the surface. Examination of early stages of biofilm
formation revealed that the marked increase in the ?flgE bio-
film occurred several hours after the initial attachment during
the maturation phase (see Fig. S3 in the supplemental mate-
rial). These nonmotile bacteria are limited to passive surface
interactions and likely colonize proximal to their site of origin,
leading to rapid accumulation of dense biofilms.
Chemotaxis influences biofilm structure under flow condi-
tions. The role of chemotaxis in bacterial biofilm formation is
unclear, with different reports concluding that chemotaxis is
either required or dispensable, depending on the model sys-
8012MERRITT ET AL.J. BACTERIOL.
tem. For example, Pratt and Kolter concluded that chemotaxis
is fully dispensable for biofilm formation in E. coli (33). In
contrast, Aeromonas sp. chemotaxis mutants have been shown
to have a severe biofilm defect in static culture (20). Collec-
tively, these observations indicate that there are species-spe-
cific differences in the requirement for chemotaxis during bio-
While the static culture biofilm deficiency manifested by the
?cheA mutant appears to be mild when corrected for culture
growth, short-term binding assays showed that this mutant is in
fact significantly impaired for surface attachment (Fig. 3B).
This defect is largely independent of growth because the short
time scale of these experiments does not allow extensive cell
division to occur. Therefore, the ?cheA mutant attachment
and biofilm phenotypes strongly suggest that there is a role for
chemotaxis, and therefore regulated motility, in surface sam-
pling, attachment, and subsequent biofilm formation. Flow cell
biofilms of the ?cheA mutant were consistently less confluent
and more structurally homogeneous than those of the wild type
(Fig. 5). The decreased density of mature ?cheA biofilms (Fig.
4D) may reflect reduced attachment efficiency as well as a role
for chemotaxis in determining the cellular organization within
the biofilm and, ultimately, its architecture. Although this phe-
notype could also be influenced by reduced growth of the
?cheA mutant, the unique structural attributes (Fig. 4 and 6) of
this biofilm strongly suggest more specific roles for chemotaxis
in biofilm formation.
A chemotaxis mutant suppressor reveals the importance of
appropriate flagellar control in biofilm formation. During the
analysis of our ?cheA mutant, we isolated the cms suppressors
of the Che?swim defect and subsequently found these mu-
tants to be profoundly debilitated for biofilm formation. The
observed restoration of tumbling in the suppressor mutants
was not due to a true reversal of the chemotaxis phenotype, as
demonstrated by the normal swim ring diameter of a cms-1
mutant expressing a plasmid-borne cheA gene (Fig. 7). Sup-
pression was also observed in other Che?mutants, including
one with a clean deletion of the entire che operon, and there-
fore it is likely that the swimming phenotype of the suppressor
functions independently of the chemotaxis system. The Cms
phenotype bears facile similarity to a swim plate phenotype
described as pseudotaxis, which is correlated with increased
tumbling in an E. coli mutant containing a deletion of the
entire che operon (44). Several E. coli and Salmonella enterica
serovar Typhimurium pseudotaxis mutations have been
mapped to the flagellar switch gene fliM (38). These FliM
mutations increase the basal frequency of rotational switching
for the flagella, resulting in a larger swim ring diameter on
motility plates (38, 44). We have not identified any mutations
in fliM (Atu0561) or several other C58 candidate genes which
could influence tumbling frequency (data not shown), although
this does not rule out the possibility that the cms mutants are
The cms-1 mutant is severely deficient in attachment and
subsequent biofilm formation under static conditions. This bio-
film phenotype was not rescued by complementation with
cheA, suggesting that the cms-1 defect is independent of che-
motaxis. It is plausible that the restored yet aberrant tumbling
exhibited by the cms-1 mutant interferes with productive sur-
face interactions. Although it is formally possible that this
mutation leads to an additional defect unrelated to motility,
this seems unlikely given the overall importance of this process
in C58 biofilm formation.
In contrast to static culture, biofilms of the cms-1 mutant
and its ?cheA parent were strikingly similar in flow cells,
though notable differences remained. Biofilms of the cms-1
mutant had greater average heights and were even less densely
packed with cells than the ?cheA biofilms (Fig. 4 and 5). The
biofilm deficiencies manifested by the cms-1 mutant, in either
a flowing or static environment, indicate that the renewed
tumbling exhibited by this mutant does not promote productive
surface interactions. The control of flagellar rotation is likely to
be critical for initial attachment and the subsequent transition
from a planktonic to a sessile state. Identification of the mu-
tation(s) underlying the Cms suppression phenotype should
provide insights into how A. tumefaciens coordinates control of
flagellar activity during the early stages of biofilm formation.
Motility and chemotaxis during the transition to sessile
growth. One reasonable model for the role of motility and
chemotaxis in biofilm formation by strain C58 is that the tran-
sition from initial to permanent attachment requires regulation
of flagellar rotation. In static culture, flagella serve to promote
surface interactions by propelling cells to the surface. The
efficiency and frequency of surface sampling may be influenced
through chemotactic processes. During the sampling period,
the flagella are still able to rotate, allowing cells to return to
the planktonic phase. Once the transition to a sessile growth
state has been made, motility is no longer required. As the
biofilm matures, other signals, such as chemotactic cues, may
stimulate renewed motility. Bacteria released into the plank-
tonic phase have the potential to colonize new sites and pro-
mote lateral expansion of the biofilm. Our findings show that
both motility and chemotaxis contribute to biofilm formation
and demonstrate the significant influence of the prevailing
environment on their respective functions. A. tumefaciens C58
differs from other biofilm model systems, such as P. aeruginosa,
in its lack of surface motility. This allows analysis of the effects
of swimming motility without the confounding processes of
twitching or swarming.
We acknowledge Michael Hibbing, Yves Brun, Dan Kearns, Tom
Platt, and Ce ´cile Berne for helpful discussions. Cherı ´e Blair provided
valuable technical support.
This project was supported by National Institutes of Health grant
GM080546 and through a grant from the Indiana University META-
Cyt program, funded in part by a major endowment from the Lilly
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