JOURNAL OF BACTERIOLOGY, Nov. 2008, p. 7392–7405
Copyright © 2008, American Society for Microbiology. All Rights Reserved.
Vol. 190, No. 22
Identification and Characterization of Cyclic Diguanylate Signaling
Systems Controlling Rugosity in Vibrio cholerae?†
Sinem Beyhan, Lindsay S. Odell, and Fitnat H. Yildiz*
Department of Microbiology and Environmental Toxicology, University of California, Santa Cruz, Santa Cruz, California 95064
Received 23 April 2008/Accepted 29 August 2008
Vibrio cholerae, the causative agent of the disease cholera, can generate rugose variants that have an
increased capacity to form biofilms. Rugosity and biofilm formation are critical for the environmental survival
and transmission of the pathogen, and these processes are controlled by cyclic diguanylate (c-di-GMP)
signaling systems. c-di-GMP is produced by diguanylate cyclases (DGCs) and degraded by phosphodiesterases
(PDEs). Proteins that contain GGDEF domains act as DGCs, whereas proteins that contain EAL or HD-GYP
domains act as PDEs. In the V. cholerae genome there are 62 genes that are predicted to encode proteins
capable of modulating the cellular c-di-GMP concentration. We previously identified two DGCs, VpvC and
CdgA, that can control the switch between smooth and rugose. To identify other c-di-GMP signaling proteins
involved in rugosity, we generated in-frame deletion mutants of all genes predicted to encode proteins with
GGDEF and EAL domains and then searched for mutants with altered rugosity. In this study, we identified two
new genes, cdgG and cdgH, involved in rugosity control. We determined that CdgH acts as a DGC and positively
regulates rugosity, whereas CdgG does not have DGC activity and negatively regulates rugosity. In addition,
epistasis analysis with CdgG, CdgH, and other DGCs and PDEs controlling rugosity revealed that CdgG and
CdgH act in parallel with previously identified c-di-GMP signaling proteins to control rugosity in V. cholerae.
We also determined that PilZ domain-containing c-di-GMP binding proteins contribute minimally to rugosity,
indicating that there are additional c-di-GMP binding proteins controlling rugosity in V. cholerae.
Vibrio cholerae causes cholera and is a natural inhabitant of
aquatic environments (14, 34). Seasonal cholera outbreaks oc-
cur where the disease is endemic and can spread worldwide. V.
cholerae’s ability to cause epidemics is tied to its ability to
survive in aquatic habitats (14, 34). One key factor that is
important for environmental survival and transmission of V.
cholerae is the microbe’s ability to form biofilms, which are
matrix-enclosed surface-associated communities (1, 14, 34).
V. cholerae can generate colony variants, termed smooth and
rugose, that differ significantly in their biofilm-forming capac-
ities (58). Smooth-to-rugose conversion occurs spontaneously
under a variety of conditions, including carbon limitation,
growth in biofilms, and treatment with bactericidal agents (39,
54, 58). Rugose variants have been isolated from environmen-
tal biofilm samples collected in Bangladesh, indicating that the
smooth-to-rugose switch can also occur in natural environ-
ments (24). In addition, Morris et al. have shown that rugose
variants can cause cholera when given orally to human volun-
teers, thus demonstrating that rugose variants can infect hu-
mans (39). Several molecular mechanisms controlling the
smooth-to-rugose switch have been found, but each of these
mechanisms does not function in all strains. The identified
molecular alterations to create rugosity include the loss of
HapR (the master regulator of quorum sensing) (19, 57, 59),
FlaA (a major flagellin subunit) (29, 55), or CytR (a regulator
of nucleoside uptake and catabolism) (20). In our prototype
strain (V. cholerae O1 El Tor, A1552), hapR mutants form
rugose colonies, but flaA and cytR mutants form smooth col-
onies. These results demonstrate that there are multiple ways
by which the smooth-to-rugose switch can take place.
Rugosity and formation of mature biofilms require extracel-
lular matrix components. A major component of the V. chol-
erae biofilm matrix is the VPS (named for Vibrio polysaccha-
ride) exopolysaccharide. VPS production is essential for the
development of three-dimensional biofilm structures (58) and
is mediated by proteins encoded by the vps genes, which are
organized into vps-I and vps-II clusters on the large chromo-
some (58). Protein components of the V. cholerae biofilm ma-
trix are also required for rugosity and the formation of a
wild-type biofilm (15, 16). Biofilm matrix production is posi-
tively controlled by transcriptional regulators VpsR and VpsT
(8, 56) and negatively regulated by the quorum-sensing tran-
scriptional regulator HapR (19, 57, 59), as well as the cyclic
AMP (cAMP) and cyclic AMP receptor protein regulatory
Cyclic diguanylate (c-di-GMP) has emerged as a ubiquitous
second messenger in bacteria that controls the transition from
a free-living, motile lifestyle to a biofilm lifestyle (42). c-di-
GMP production and degradation is controlled by diguanylate
cyclases (DGCs) and phosphodiesterases (PDEs), respectively.
Proteins that contain GGDEF domains act as DGCs, whereas
proteins that contain EAL or HD-GYP domains act as PDEs
(44, 46, 47). In addition, proteins carrying PilZ domains, which
are shown to bind c-di-GMP, are one type of downstream
protein that relays signals to cellular processes (11, 36, 40, 45).
V. cholerae has 31 genes that encode proteins with a GGDEF
domain, 12 genes that encode proteins with an EAL domain,
* Corresponding author. Mailing address: Department of Microbi-
ology and Environmental Toxicology, University of California, Santa
Cruz, Santa Cruz, CA 95064. Phone: (831) 459-1588. Fax: (831) 459-
3524. E-mail: firstname.lastname@example.org.
† Supplemental material for this article may be found at http://jb
?Published ahead of print on 12 September 2008.
10 genes that encodes proteins with both GGDEF and EAL
domains, 9 genes that encode proteins with a HD-GYP do-
main, and 4 genes that encode proteins with a conserved PilZ
domain (2, 18). Studies to date have shown that c-di-GMP
regulates biofilm formation, motility, virulence, and smooth-
to-rugose phase variation in V. cholerae (5–7, 27, 32, 40, 41, 50,
52, 53). We recently demonstrated that rugose variants have
increased c-di-GMP levels compared to smooth variants, lead-
ing to elevated biofilm formation in the rugose forms (4, 6, 32).
The rugosity-associated increase in c-di-GMP in our prototype
rugose strain is caused by a single amino acid change in a DGC
protein, which we called VpvC (6). However, other genetic
variations can also cause rugosity. For example, we found that
increased transcription of cdgA, encoding another DGC,
causes rugosity in hapR mutants (4). Furthermore, an increase
in c-di-GMP due to loss of a key PDE, CdgC, MbaA, or RocS,
in the rugose genetic background leads to formation of super-
rugose colonies that are more opaque and corrugated than
rugose (7, 32, 41).
In the present study, we investigated whether other genes
encoding DGCs and PDEs contribute to rugosity and, in turn,
biofilm formation in V. cholerae. We identified and character-
ized two such genes, cdgG and cdgH, encoding GGDEF do-
main proteins. Through epistasis analysis, we determined that
many c-di-GMP signaling proteins act in parallel pathways to
control rugosity. We also determined that PilZ domain-con-
taining c-di-GMP receptors contribute minimally to rugosity,
indicating that there are additional c-di-GMP receptors con-
MATERIALS AND METHODS
Bacterial strains and growth conditions. Bacterial strains and plasmids used in
the present study are listed in Table 1. Escherichia coli DH10B and CC118 ?pir
strains were used for DNA manipulation, and E. coli S17-1 ?pir strains were used
for conjugation with V. cholerae. Knockout mutants of V. cholerae strains and V.
cholerae strains carrying plasmids with lacZ transcriptional fusions and multicopy
vectors were generated as described earlier (32). V. cholerae cultures were grown
in Luria-Bertani (LB) broth (1% tryptone, 0.5% yeast extract, 1% NaCl [pH 7.5])
with aeration at 30°C. E. coli cultures were grown in the same medium with
aeration at 37°C. Antibiotics (rifampin and ampicillin) were added at 100 ?g/ml
unless otherwise noted. For the induction of gene expression in strains carrying
arabinose-inducible vectors, L-arabinose was added to the growth medium at a
final concentration of 0.2% (wt/vol) for c-di-GMP quantification experiments
and 0.02% (wt/vol) for complementation analysis via colony morphology and
DNA manipulations. The DNA oligonucleotides used in the present study
were purchased from Operon Technologies (Alameda, CA) and are listed in
Table S1 in the supplemental material. A PCR method was used to generate
in-frame deletions of the cdgG and cdgH genes utilizing previously published
methods (32). Deletion vectors were constructed by using pGP704sac28 suicide
plasmid as described previously (32). For overexpression studies, cdgG and cdgH
were cloned into pBAD/myc-His-B plasmid using the primers listed (see Table
S1 in the supplemental material). Construction of cdgG and cdgH vectors har-
boring point mutations was done utilizing a previously published method (6). The
deletion and overexpression constructs were sequenced (UC Berkeley DNA
Sequencing Facility), and only the clones without any undesired mutations were
Generation of V. cholerae deletion mutants and green fluorescent protein
tagging of V. cholerae strains. V. cholerae deletion mutants and green fluorescent
protein-tagged V. cholerae strains were generated as described previously
?-Galactosidase assays. ?-Galactosidase assays were performed and Miller
units were calculated as previously described (32, 37).
c-di-GMP quantification. The amount of c-di-GMP was quantified by two-
dimensional thin-layer chromatography (2D-TLC) as previously described (32,
52) with the following modifications. Briefly, bacteria were grown in morpho-
linepropanesulfonic acid (MOPS) minimal medium containing 0.75 mM
KH2PO4at 30°C for overnight. The cells were then diluted 1:50 into fresh MOPS
minimal medium containing 0.25 mM KH2PO4and grown to an optical density
at 600 nm of 0.6 ? 0.05 at 30°C. Then, 50 ?Ci of [32P]orthophosphate was added
to 0.5 ml of cell suspension, followed by further growth for 1 h. Labeled nucle-
otides were extracted by using previously published methods (52). Portions (10
?l) of total nucleotides were separated on TLC plates as previously described
Colony morphology. For colony morphology assays, V. cholerae colonies were
grown on LB agar plates for 1 to 2 days at 30°C. Colonies were photographed by
using a Nikon CoolPix 4500 digital camera.
Biofilm assays. The biofilm-forming capacities of the V. cholerae strains were
determined using cover glass chambers (Lab-Tek). Dilutions (1:100, 2 ml) in LB
medium from overnight-grown cultures were placed into chambers. Biofilms
were formed under static conditions at 30°C for 8 h, washed twice with 1 ml of
LB medium, and then visualized by using confocal laser scanning microscopy
(CLSM). Acquired images were analyzed with the COMSTAT program (22). For
flow cell experiments, biofilms were grown at room temperature in flow cham-
bers (individual channel dimensions of 1 by 4 by 40 mm) supplied with 2% LB
medium supplemented with 0.9% NaCl (0.02% peptone, 0.01% yeast extract,
0.9% NaCl) at a flow rate of 4.5 ml/h. Assembly of the flow cell system and image
acquisitions were done as previously described (57).
Motility assays. LB soft agar plates (0.3% agar) were used to determine the
motility of bacterial strains (57). The diameter of the migration zone was mea-
sured after 18 h of incubation at 30°C.
CdgH positively regulates rugosity-associated phenotypes.
In V. cholerae, there are a total of 62 genes that are predicted
to encode proteins capable of producing or degrading c-di-
GMP (18). Fifty-three of these genes encode proteins with
GGDEF and/or EAL domains. We previously studied 10 of
them and had shown that VpvC, CdgA, CdgC, MbaA, and
RocS regulate rugosity in V. cholerae O1 El Tor A1552 strain
(4, 6, 32). In the present study, we analyzed the contribution of
the remaining genes encoding proteins with GGDEF and/or
EAL domains to rugosity. To this end, we created in-frame
deletions of all GGDEF/EAL genes (except VC0515, since the
genomic region appears to be different between our prototype
strain and the sequenced N16961 strain) in the rugose genetic
background; we then screened the mutants for alterations in
colony corrugation when grown on LB agar plates. Through
this screen of 42 mutants, we identified two additional mutants
exhibiting a significant change in colony corrugation.
The first mutant had a deletion in VC1067, which encodes a
protein with a GGDEF domain, now termed cdgH for cyclic
diguanylate H. The second mutant had a deletion in VC0900,
which encodes a protein with a GGDEF domain, now termed
cdgG for cyclic diguanylate G.
We first investigated the contribution of CdgH to rugosity-
associated phenotypes by analyzing colony corrugation and
biofilm formation. Colonies formed by the rugose cdgH mutant
(R?cdgH) on LB agar plates appeared less corrugated and
flatter compared to those formed by rugose, a finding consis-
tent with producing less VPS (Fig. 1A). Similarly, R?cdgH
formed thinner and less-structured biofilms compared to ru-
gose after 8 h of biofilm development at 30°C under static
conditions (Fig. 1B). To quantify the differences in biofilm
architecture, we used the COMSTAT program and quantified
various biofilm parameters: biomass, average thickness, maxi-
mum thickness, substratum coverage, roughness, and the sur-
face area/volume ratio. Although, substratum coverage, rough-
ness, and the surface area/volume ratio remained the same in
VOL. 190, 2008EFFECT OF CdgG AND CdgH ON RUGOSITY 7393
affecting mainly protein activity (43), recent studies indicate
that c-di-GMP can regulate gene expression by interacting with
transcriptional factors (23) and through cyclic di-GMP ribo-
switches (48). We have studied c-di-GMP signaling proteins
that affect colony rugosity. Several of these alter gene expres-
sion, including VpvC, CdgA, CdgH, CdgC, RocS, and MbaA
(4, 6, 33), while CdgG apparently does not. These findings
indicate that both transcriptional and posttranscriptional
modes of regulation operate in V. cholerae c-di-GMP signaling
systems. V. cholerae has 62 genes predicted to encode proteins
with GGDEF, EAL, or HD-GYP domains and faces another
challenge, namely, how an organism with a large number of
c-di-GMP signaling proteins can prevent cross talk or noise in
signaling. Spatial sequestration of GGDEF/EAL proteins to
microcompartmentalize c-di-GMP levels in the cell is one of
the proposed mechanisms. VpvC, CdgA, and CdgH are pre-
dicted to be localized to the cytoplasmic membrane. We pro-
pose that they could form independent c-di-GMP signaling
clusters in different regions of the cell, together with their
effector proteins (transcriptional regulators or proteins that
can change activity or function of transcriptional regulators). It
is likely that each of these proteins generate a different c-di-
GMP pool that can be degraded by cognate PDEs (Fig. 12).
c-di-GMP signaling complexes could be dynamic and effector
proteins could be released from these complexes and partici-
pate directly or indirectly in gene expression. Alternatively,
some portion of c-di-GMP may freely diffuse in the cytoplasm,
and c-di-GMP might interact with cytoplasmically localized
transcriptional regulators, thereby regulating gene expression.
We undertook systematic mutational and phenotypic analyses
of c-di-GMP signaling proteins in V. cholerae and identified
critical c-di-GMP signaling proteins required for rugosity-as-
sociated phenotypes. There is much to be discovered about the
mechanisms by which the c-di-GMP signaling system regulates
rugosity and interacts with other regulatory networks control-
ling rugosity, including two component regulatory systems and
quorum sensing. A better understanding of the mechanism of
c-di-GMP signaling and biofilm formation and the importance
of these processes in V. cholerae biology will prove useful for
the development of future strategies for predicting and con-
trolling cholera epidemics.
We thank Bentley Lim for constructing pFY-572; Vanessa Soliven
for generating Fy_Vc_337 and Fy_Vc_339; and Karen Ottemann,
Manel Camps, and Yildiz laboratory members for their valuable com-
ments on the manuscript.
This study was supported by NIH grant AI055987.
1. Alam, M., M. Sultana, G. B. Nair, R. B. Sack, D. A. Sack, A. K. Siddique, A.
Ali, A. Huq, and R. R. Colwell. 2006. Toxigenic Vibrio cholerae in the aquatic
environment of Mathbaria, Bangladesh. Appl. Environ. Microbiol. 72:2849–
2. Amikam, D., and M. Y. Galperin. 2006. PilZ domain is part of the bacterial
c-di-GMP binding protein. Bioinformatics 22:3–6.
3. Bao, Y., D. P. Lies, H. Fu, and G. P. Roberts. 1991. An improved Tn7-based
system for the single-copy insertion of cloned genes into chromosomes of
gram-negative bacteria. Gene 109:167–168.
4. Beyhan, S., K. Bilecen, S. R. Salama, C. Casper-Lindley, and F. H. Yildiz.
2007. Regulation of rugosity and biofilm formation in Vibrio cholerae: com-
parison of VpsT and VpsR regulons and epistasis analysis of vpsT, vpsR, and
hapR. J. Bacteriol. 189:388–402.
5. Beyhan, S., A. D. Tischler, A. Camilli, and F. H. Yildiz. 2006. Transcriptome
and phenotypic responses of Vibrio cholerae to increased cyclic di-GMP level.
J. Bacteriol. 188:3600–3613.
6. Beyhan, S., and F. H. Yildiz. 2007. Smooth to rugose phase variation in
Vibrio cholerae can be mediated by a single nucleotide change that targets
c-di-GMP signalling pathway. Mol. Microbiol. 63:995–1007.
7. Bomchil, N., P. Watnick, and R. Kolter. 2003. Identification and character-
ization of a Vibrio cholerae gene, mbaA, involved in maintenance of biofilm
architecture. J. Bacteriol. 185:1384–1390.
8. Casper-Lindley, C., and F. H. Yildiz. 2004. VpsT is a transcriptional regu-
lator required for expression of vps biosynthesis genes and the development
of rugose colonial morphology in Vibrio cholerae O1 El Tor. J. Bacteriol.
9. Chan, C., R. Paul, D. Samoray, N. C. Amiot, B. Giese, U. Jenal, and T.
Schirmer. 2004. Structural basis of activity and allosteric control of digua-
nylate cyclase. Proc. Natl. Acad. Sci. USA 101:17084–17089.
10. Christen, B., M. Christen, R. Paul, F. Schmid, M. Folcher, P. Jenoe, M.
Meuwly, and U. Jenal. 2006. Allosteric control of cyclic di-GMP signaling.
J. Biol. Chem. 104:4112–4117.
11. Christen, M., B. Christen, M. G. Allan, M. Folcher, P. Jeno, S. Grzesiek, and
U. Jenal. 2007. DgrA is a member of a new family of cyclic diguanosine
monophosphate receptors and controls flagellar motor function in
Caulobacter crescentus. Proc. Natl. Acad. Sci. USA 104:4112–4117.
12. Christen, M., B. Christen, M. Folcher, A. Schauerte, and U. Jenal. 2005.
Identification and characterization of a cyclic di-GMP-specific phosphodi-
esterase and its allosteric control by GTP. J. Biol. Chem. 280:30829–30837.
13. de Lorenzo, V., and K. N. Timmis. 1994. Analysis and construction of stable
phenotypes in gram-negative bacteria with Tn5- and Tn10-derived minitrans-
posons. Methods Enzymol. 235:386–405.
14. Faruque, S. M., M. J. Albert, and J. J. Mekalanos. 1998. Epidemiology,
genetics, and ecology of toxigenic Vibrio cholerae. Microbiol. Mol. Biol. Rev.
15. Fong, J. C., K. Karplus, G. K. Schoolnik, and F. H. Yildiz. 2006. Identifica-
tion and characterization of RbmA, a novel protein required for the devel-
opment of rugose colony morphology and biofilm structure in Vibrio chol-
erae. J. Bacteriol. 188:1049–1059.
16. Fong, J. C., and F. H. Yildiz. 2007. The rbmBCDEF gene cluster modulates
development of rugose colony morphology and biofilm formation in Vibrio
cholerae. J. Bacteriol. 189:2319–2330.
17. Fullner, K. J., and J. J. Mekalanos. 1999. Genetic characterization of a new
type IV-A pilus gene cluster found in both classical and El Tor biotypes of
Vibrio cholerae. Infect. Immun. 67:1393–1404.
18. Galperin, M. Y., A. N. Nikolskaya, and E. V. Koonin. 2001. Novel domains
of the prokaryotic two-component signal transduction systems. FEMS Mi-
crobiol. Lett. 203:11–21.
19. Hammer, B. K., and B. L. Bassler. 2003. Quorum sensing controls biofilm
formation in Vibrio cholerae. Mol. Microbiol. 50:101–104.
20. Haugo, A. J., and P. I. Watnick. 2002. Vibrio cholerae CytR is a repressor of
biofilm development. Mol. Microbiol. 45:471–483.
21. Herrero, M., V. de Lorenzo, and K. N. Timmis. 1990. Transposon vectors
containing non-antibiotic resistance selection markers for cloning and stable
chromosomal insertion of foreign genes in gram-negative bacteria. J. Bacte-
22. Heydorn, A., A. T. Nielsen, M. Hentzer, C. Sternberg, M. Givskov, B. K.
Ersboll, and S. Molin. 2000. Quantification of biofilm structures by the novel
computer program COMSTAT. Microbiology 146(Pt. 10):2395–2407.
23. Hickman, J. W., and C. S. Harwood. 2008. Identification of FleQ from
Pseudomonas aeruginosa as a c-di-GMP-responsive transcription factor. Mol.
24. Islam, M. S., M. I. Jahid, M. M. Rahman, M. Z. Rahman, M. S. Islam, M. S.
Kabir, D. A. Sack, and G. K. Schoolnik. 2007. Biofilm acts as a microenvi-
ronment for plankton-associated Vibrio cholerae in the aquatic environment
of Bangladesh. Microbiol. Immunol. 51:369–379.
25. Karatan, E., T. R. Duncan, and P. I. Watnick. 2005. NspS, a predicted
polyamine sensor, mediates activation of Vibrio cholerae biofilm formation by
norspermidine. J. Bacteriol. 187:7434–7443.
26. Kazmierczak, B. I., M. B. Lebron, and T. S. Murray. 2006. Analysis of FimX,
a phosphodiesterase that governs twitching motility in Pseudomonas aerugi-
nosa. Mol. Microbiol. 60:1026–1043.
27. Kovacikova, G., W. Lin, and K. Skorupski. 2005. Dual regulation of genes
involved in acetoin biosynthesis and motility/biofilm formation by the viru-
lence activator AphA and the acetate-responsive LysR-type regulator AlsR
in Vibrio cholerae. Mol. Microbiol. 57:420–433.
28. Kuchma, S. L., K. M. Brothers, J. H. Merritt, N. T. Liberati, F. M. Ausubel,
and G. A. O’Toole. 2007. BifA, a cyclic-Di-GMP phosphodiesterase, in-
versely regulates biofilm formation and swarming motility by Pseudomonas
aeruginosa PA14. J. Bacteriol. 189:8165–8178.
29. Lauriano, C. M., C. Ghosh, N. E. Correa, and K. E. Klose. 2004. The
sodium-driven flagellar motor controls exopolysaccharide expression in
Vibrio cholerae. J. Bacteriol. 186:4864–4874.
30. Lee, V. T., J. M. Matewish, J. L. Kessler, M. Hyodo, Y. Hayakawa, and S.
Lory. 2007. A cyclic-di-GMP receptor required for bacterial exopolysaccha-
ride production. Mol. Microbiol. 65:1474–1484.
7404 BEYHAN ET AL. J. BACTERIOL.
31. Liang, W., A. J. Silva, and J. A. Benitez. 2007. The cyclic AMP receptor
protein modulates colonial morphology in Vibrio cholerae. Appl. Environ.
32. Lim, B., S. Beyhan, J. Meir, and F. H. Yildiz. 2006. Cyclic-diGMP signal
transduction systems in Vibrio cholerae: modulation of rugosity and biofilm
formation. Mol. Microbiol. 60:331–348.
33. Lim, B., S. Beyhan, and F. H. Yildiz. 2007. Regulation of Vibrio polysaccha-
ride synthesis and virulence factor production by CdgC, a GGDEF-EAL
domain protein, in Vibrio cholerae. J. Bacteriol. 189:717–729.
34. Lipp, E. K., A. Huq, and R. R. Colwell. 2002. Effects of global climate on
infectious disease: the cholera model. Clin. Microbiol. Rev. 15:757–770.
35. Malone, J. G., R. Williams, M. Christen, U. Jenal, A. J. Spiers, and P. B.
Rainey. 2007. The structure-function relationship of WspR, a Pseudomonas
fluorescens response regulator with a GGDEF output domain. Microbiology
36. Merighi, M., V. T. Lee, M. Hyodo, Y. Hayakawa, and S. Lory. 2007. The
second messenger bis-(3?-5?)-cyclic-GMP and its PilZ domain-containing
receptor Alg44 are required for alginate biosynthesis in Pseudomonas aerugi-
nosa. Mol. Microbiol. 65:876–895.
37. Miller, J. H. 1972. Assay of ?-galactosidase. Cold Spring Harbor Laboratory,
Cold Spring Harbor, NY.
38. Moorthy, S., and P. I. Watnick. 2004. Genetic evidence that the Vibrio
cholerae monolayer is a distinct stage in biofilm development. Mol. Micro-
39. Morris, J. G., Jr., M. B. Sztein, E. W. Rice, J. P. Nataro, G. A. Losonsky, P.
Panigrahi, C. O. Tacket, and J. A. Johnson. 1996. Vibrio cholerae O1 can
assume a chlorine-resistant rugose survival form that is virulent for humans.
J. Infect. Dis. 174:1364–1368.
40. Pratt, J. T., R. Tamayo, A. D. Tischler, and A. Camilli. 2007. PilZ domain
proteins bind cyclic diguanylate and regulate diverse processes in Vibrio
cholerae. J. Biol. Chem. 282:12860–12870.
41. Rashid, M. H., C. Rajanna, A. Ali, and D. K. Karaolis. 2003. Identification
of genes involved in the switch between the smooth and rugose phenotypes
of Vibrio cholerae. FEMS Microbiol. Lett. 227:113–119.
42. Romling, U., and D. Amikam. 2006. Cyclic di-GMP as a second messenger.
Curr. Opin. Microbiol. 9:218–228.
43. Ross, P., H. Weinhouse, Y. Aloni, D. Michaeli, P. Weinbergerohana, R.
Mayer, S. Braun, E. Devroom, G. Vandermarel, J. Vanboom, and M. Ben-
ziman. 1987. Regulation of cellulose synthesis in Acetobacter-xylinum by
cyclic digualylic acid. Nature 325:279–281.
44. Ryan, R. P., Y. Fouhy, J. F. Lucey, L. C. Crossman, S. Spiro, Y. W. He, L. H.
Zhang, S. Heeb, M. Camara, P. Williams, and J. M. Dow. 2006. Cell-cell
signaling in Xanthomonas campestris involves an HD-GYP domain protein
that functions in cyclic di-GMP turnover. Proc. Natl. Acad. Sci. USA 103:
45. Ryjenkov, D. A., R. Simm, U. Romling, and M. Gomelsky. 2006. The PilZ
domain is a receptor for the second messenger c-di-GMP: the PilZ domain
protein YcgR controls motility in enterobacteria. J. Biol. Chem. 281:30310–
46. Ryjenkov, D. A., M. Tarutina, O. V. Moskvin, and M. Gomelsky. 2005. Cyclic
diguanylate is a ubiquitous signaling molecule in bacteria: insights into bio-
chemistry of the GGDEF protein domain. J. Bacteriol. 187:1792–1798.
47. Schmidt, A. J., D. A. Ryjenkov, and M. Gomelsky. 2005. The ubiquitous
protein domain EAL is a cyclic diguanylate-specific phosphodiesterase: en-
zymatically active and inactive EAL domains. J. Bacteriol. 187:4774–4781.
48. Sudarsan, N., E. R. Lee, Z. Weinberg, R. H. Moy, J. N. Kim, K. H. Link, and
R. R. Breaker. 2008. Riboswitches in eubacteria sense the second messenger
cyclic di-GMP. Science 321:411–413.
49. Suzuki, K., P. Babitzke, S. R. Kushner, and T. Romeo. 2006. Identification
of a novel regulatory protein (CsrD) that targets the global regulatory RNAs
CsrB and CsrC for degradation by RNase E. Genes Dev. 20:2605–2617.
50. Tamayo, R., S. Schild, J. T. Pratt, and A. Camilli. 2008. Role of cyclic
di-GMP during El Tor biotype Vibrio cholerae infection: characterization of
the in vivo-induced cyclic di-GMP phosphodiesterase CdpA. Infect. Immun.
51. Thormann, K. M., S. Duttler, R. M. Saville, M. Hyodo, S. Shukla, Y. Hay-
akawa, and A. M. Spormann. 2006. Control of formation and cellular de-
tachment from Shewanella oneidensis MR-1 biofilms by cyclic di-GMP. J.
52. Tischler, A. D., and A. Camilli. 2004. Cyclic diguanylate (c-di-GMP) regu-
lates Vibrio cholerae biofilm formation. Mol. Microbiol. 53:857–869.
53. Tischler, A. D., and A. Camilli. 2005. Cyclic diguanylate regulates Vibrio
cholerae virulence gene expression. Infect. Immun. 73:5873–5882.
54. Wai, S. N., Y. Mizunoe, A. Takade, S. I. Kawabata, and S. I. Yoshida. 1998.
Vibrio cholerae O1 strain TSI-4 produces the exopolysaccharide materials
that determine colony morphology, stress resistance, and biofilm formation.
Appl. Environ. Microbiol. 64:3648–3655.
55. Watnick, P. I., C. M. Lauriano, K. E. Klose, L. Croal, and R. Kolter. 2001.
The absence of a flagellum leads to altered colony morphology, biofilm
development and virulence in Vibrio cholerae O139. Mol. Microbiol. 39:223–
56. Yildiz, F. H., N. A. Dolganov, and G. K. Schoolnik. 2001. VpsR, a member
of the response regulators of the two-component regulatory systems, is
required for expression of vps biosynthesis genes and EPS(ETr)-associated
phenotypes in Vibrio cholerae O1 El Tor. J. Bacteriol. 183:1716–1726.
57. Yildiz, F. H., X. S. Liu, A. Heydorn, and G. K. Schoolnik. 2004. Molecular
analysis of rugosity in a Vibrio cholerae O1 El Tor phase variant. Mol.
58. Yildiz, F. H., and G. K. Schoolnik. 1999. Vibrio cholerae O1 El Tor: identi-
fication of a gene cluster required for the rugose colony type, exopolysac-
charide production, chlorine resistance, and biofilm formation. Proc. Natl.
Acad. Sci. USA 96:4028–4033.
59. Zhu, J., and J. J. Mekalanos. 2003. Quorum sensing-dependent biofilms
enhance colonization in Vibrio cholerae. Dev. Cell 5:647–656.
VOL. 190, 2008EFFECT OF CdgG AND CdgH ON RUGOSITY 7405