MOLECULAR BIOLOGY AND BIOCHEMISTRY OF THE
REGULATION OF HRP/TYPE III SECRETION GENES IN THE
CORN PATHOGEN PANTOEA STEWARTII SUBSP. STEWARTII
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the
Degree Doctor of Philosophy in the
Graduate School of The Ohio State University
By
Massimo Merighi, B. S., M. S.
*****
The Ohio State University
2003
Dissertation Committee:
Professor David L. Coplin, Adviser Approved by
Professor Brian Ahmer ______
Professor Dietz W. Bauer Adviser
Professor Terrence L. Graham Plant Pathology
Copyright by
Massimo Merighi
2003
ii
ABSTRACT
Pantoea stewartii subsp. stewartii is a bacterial pathogen of corn. Its
pathogenicity depends on the expression of a Hrp/type III protein
secretion/translocation system. The regulatory region of the hrp gene cluster consists of three adjacent operons: hrpXY encodes a two- component regulatory system, consisting of the response regulator HrpY and sensor PAS-kinase HrpX; hrpS encodes an NtrC-like enhancer-binding protein; and hrpL encodes an ECF sigma factor. In this study, we used genetic and biochemical approaches to delineate the following regulatory cascade: 1) HrpY activates hrpS; 2) HrpS activates hrpL; and 3) HrpL activates secretion and effector genes with ‘Hrp-box’ promoters. This pathway responds to environmental signals and global regulators.
Mutant analysis showed that HrpX is required for full virulence.
Deletion of its individual PAS-sensory domains revealed that they are not redundant and each may have a different role in modulating kinase activity. HrpX probably senses an intracellular signal, possibly related to nitrogen metabolism. pH, osmolarity and nicotinic acid controlled expression of hrpS independently of HrpX/HrpY.
hrpY mutants were completely avirulent. Site-directed mutagenesis of the conserved D57 residue in the HrpY receiver domain showed that it is
ii required for virulence and activation of hrpS and hrpXY in vivo. In vitro studies using purified His6 -HrpY indicated that D57 is the only
phosphorylation site and that His6 -HrpY~P has increased affinity for
PhrpS. Primer extension and deletion analyses located the PhrpS upstream of
an IS-like element and showed that transcription of hrpL from a σ 54 promoter requires an upstream activating sequence. We also discovered that the absence of transcriptional terminators downstream of hrpL and hrpY creates a novel autoregulatory loop, wherein activation of hrpL by
HrpS results in read-through transcription into hrpY and hrpS. Finally, the
EsaR/EsaI quorum sensing system was shown to modulate the expression of hrp genes and virulence. Expression of hrpS was reduced in an esaI background and restored in an esaIR strain.
iii
“...Fatti non foste per viver come bruti,
ma per seguir virtute e canoscenza’.”
(“…(Men) were not created to live like wild animals,
but to pursue virtue and knowledge.”)
(Dante’s Inferno, Canto XXVI, 119-120)
Dedicated to my parents, Graziella and Camillo
iv
ACKNOWLEDGMENTS
First of all, I would like to thank my family, dad, mom, and sister, for
their unconditional support and love throughout all these years so far
away from my motherland Italy. I know how much sacrifice that has
meant for you and I hope I made you proud in the end. Secondly, thanks
to my lovely wife, Serena. Simply put, nothing of this could have ever
happened without you. I will never forget your patience and
encouragement during these last six years we have been living together
here in America. Serena also helped me with many of the figures in
Chapter 1.
Obviously, I want to thank Dr. David Coplin, my adviser, for his
mentorship, and the intellectual freedom and support I have enjoyed in the
years spent in his lab. There I fully discovered my vocation as a
molecular bacteriologist and I found the courage to explore many new
areas of research. Secondly, I also have to thank him for the extensive
editing of this dissertation. I also want to express my gratitude to the
members of my SAC, Dr. Brian Ahmer, Dr. Dietz W. Bauer, and Dr.
Terrence L. Graham, for many stimulating discussions and for their
unconditional availability whenever I bugged them for using equipment or
v discussing ideas. In particular, a special thanks goes to Dr. Bauer, for teaching me many things about academic life and beyond, and for trying, unsuccessfully, to make a decent pool player out of me (Sorry Dietz!).
A special acknowledgement goes to current members of the lab, Doris
R. Majerczak and Dr. Jong Hyun Ham. Doris has been a good friend and a wonderful co-worker, always available and extremely helpful. Many of the experiments in this dissertation are a result of the strict cooperation between the two of us. I will always carry with me a good memory of the time spent in the lab together. Jong Hyun has been a good friend with whom I shared many nights in the lab and many discussions about science and our futures.
A particular thanks goes to Dr. Max Teplitski, who has been a dear friend of my wife and me. He has been very close to us both in times of bad and good luck and we will never forget that. He also provided me with samples of purified Salmonella enterica BarA198 before publication for the experiments described in Chapter 4.
Many other friends have helped me to endure the hardships of the
Graduate School, some by discussing science with me, others by giving me different perspectives on the facts of life, some others just by being there when I needed it the most. My dear friends, Maria, Mensheng,
Ricardo, and Dr. Trudy Torto, I will never forget you all and I hope the
Fate will bring great happiness to your lives.
vi I thank also the Ohio Agricultural Research Development Center for
supporting me with a Director’s Fellowship for the first three years, the
Graduate School of the Ohio State University for granting me a
Presidential Fellowship and the Department of Plant Pathology for its support over one year and a half of my Ph.D. program.
Some of the work presented in this dissertation was completed with the support of many people at various levels. In particular, most of the data in
Chapter 2 was published in a peer-reviewed journal before the defense of
this dissertation (Merighi et al., 2003, Mol. Plant Microbe Interact 16:
238-248). For that part, I acknowledge practical contributions from E. H.
Stover (limited to the construction of strains DM701, DM729 and DM733
carrying the plasmid pRF205), who also provided the initial observations
on the linear model of regulation and on the hrpS autoregulation several
years before I came in the lab; N. Smith, for the data in Fig. 2.7; D. R.
Majerczak, for her help in performing replicates of enzyme assays, for her
independent construction of strains DM786 and plasmids pDM1296,
pDM2560 and for her practical support in various steps leading to the
construction of several plasmids and strains used in that work. I also
acknowledge intellectual contributions by D. L. Coplin in the Discussion
section of Chapter 2. Furthermore I have to acknowledge also the
contribution by D. R. Majerczak of the two Ω interposon mutants used in
Chapter 3, which Doris constructed under my supervision, and her help in
running replicates of the enzyme assays described in Chapters 2 and 6. I
vii finally thank Dr. Jyang Chuyn Jang for his friendship, mentorship and for having me in his lab as radiation user for all the experiments in Chapters
3, 4 and 5 involving radioactive isotopes.
viii
VITA
1971 Born, Italy, European Union
1990-1996: B.S. (“Laurea”), Department of Plant Pathology,
University of Bologna, Italy, Graduated 110/110,
Summa cum Laude. Experimental thesis: “Genetic
fingerprinting by PCR and PFGE of Erwinia amylovora
strains from the Mediterranean basin”. Field of Study:
Plant Pathology, Molecular Phytobacteriology.
Summer 1996: Research Intern, Max Plank Institute für Zellbiologie,
Klaus Geider’s Bacterial Molecular Genetics
Laboratory, Heidelberg, Germany.
1997-1998: Research Assistant (MIRAAF/National Plan of Plant
Biotechnology), Institute of Plant Pathology,
University of Bologna, Italy.
2001: M.S.(non-thesis), The Ohio State University
1998-present: Graduate Student, Graduate School of The Ohio State
University, Department of Plant Pathology and Plant
Molecular Biology/Biotechnology Program:
-O.A.R.D.C. Graduate Fellow (1998-2001);
ix -Graduate Research Associate (2001-2002 and 2003-
present);
-Presidential Fellow of the Ohio State University
(2002-2003)
PUBLICATIONS
Refereed publications
1. Teplitski, M., Chen, H., Rajamani, S., Gao, M., Merighi, M., Sayre,
R. T., Robinson, J. B., Rolfe, B. G., Bauer, W. D. (2004). Chlamydomonas
reinhardtii secretes compounds that mimic bacterial signals and interfere
with quorum sensing regulation in bacteria. Plant Physiol. 134:1-10.
2. Merighi, M., Majerczak, D. R., Stover, H. E. and Coplin, D. L.
(2003). The HrpX/HrpY two-component system activates hrpS expression,
the first step in the regulatory cascade controlling the Hrp regulon in
Pantoea stewartii subsp. stewartii. Mol. Plant-Microbe Interact. 16:238-
248.
3. Merighi, M., Sandrini, A., Landini, S., Girotti, S., Ghini, S.,
Malaguti, S., Bazzi, C. (2000). Chemiluminescent and colorimetric
x detection of Erwinia amylovora by immunoenzymatic determination of
PCR amplicons from plasmid pEA29. Plant Dis. 84:49-54.
4. Zhang, Y., Merighi, M., Bazzi, C., K. Geider (1998). Genomic analysis by pulsed-field gel electrophoresis of Erwinia amylovora strains from the Mediterranean region including Italy. J. Plant Path. 80:225-232.
Book Chapters
1. Merighi, M., Majerczak, D. R., and Coplin, D. L. “The hrp genes of
Pantoea stewartii are regulated by a complex system that senses environmental signals”. In: “Plant Pathogenic Bacteria” p. 201-204, S. H.
De Boer Editor, Kluwer Academic Publishers, The Netherlands, 2001.
2. Merighi, M., Sandrini, A., Landini, S., Malaguti, S., Porrini, C.,
Sabatini, A.G., Girotti, S., Ghini, S., and Bazzi, C. "Automated detection of the plant pathogen Erwinia amylovora by chemiluminescent immunoenzymatic determination of PCR amplicons from pEA29: applications to diagnosis and epidemiology". In: "Bioluminescence and
Chemiluminescence: Perspectives for the 21st Century" p. 504-507, A.
Roda, M. Pazzagli, L. J. Kricka and P. E. Stanley Editors, Wiley and
Sons, Chichester, UK, 1998.
xi
FIELD OF STUDY
Major Field: Plant Pathology
Graduate Specialization: Plant Molecular Biology and Biotechnology
Areas of Interest: Bacterial pathogenesis and regulation of virulence;
Bacterial molecular genetics and molecular biology; Biochemistry of two- component systems.
xii
TABLE OF CONTENTS
Abstract…………………………………………………………………………..ii
Dedication……………………………………………………………………….iv
Acknowledgments……………………………………………………………….v
Vita……………………………………………………………………………….ix
List of Tables…………………………………………………………………xxii
List of Figures…………………………………………………………….…xxiii
List of Abbreviations and Symbols………………………………………..xxix
Chapters:
1. LITERATURE REVIEW...... 1
1.1 Stewart’s wilt of corn and its causal agent, Pantoea stewartii subsp. stewartii ...... 1
1.1.1 Historical background ...... 1
1.1.2 Host range and resistance ...... 4
1.1.3 Geographic distribution of the disease ...... 7
1.1.4 Symptoms and signs...... 10
1.1.5 Source of inoculum, disease cycle and histopathology...... 12
1.1.6 Taxonomy ...... 16
1.1.7 Molecular Diagnostics...... 21
1.1.8 Genetics...... 23
xiii 1.1.9 Molecular pathogenesis ...... 28
1.2 Type III secretion systems, hrp genes and their regulation ...... 34
1.2.1 Bacterial secretion systems and the type III secretion apparatus. 34
1.2.2. Phytobacterial Hrp type III secretion systems...... 42
1.2.3 Regulation of type III secretion systems ...... 54
1.2.3.1 Regulation of group I hrp/hrc genes ...... 56
1.2.3.2 Regulation of group II hrp/hrc genes ...... 61
1.3 Mechanisms and strategies of two-component system regulation ..... 63
1.3.1 Regulation of virulence factors ...... 63
1.3.2 Two-component systems...... 65
1.3.2.1 Response regulators ...... 68
1.3.2.2 Sensor histidine kinases...... 70
1.4. Significance, rationale and objectives ...... 75
1.4.1 Significance ...... 75
1.4.2 Rationale ...... 77
1.4.2 Objectives...... 79
2. THE HRPX/HRPY TWO-COMPONENT SYSTEM ACTIVATES
HRPS EXPRESSION, FIRST STEP IN THE REGULATORY
CASCADE CONTROLLING THE HRP REGULON IN PANTOEA
STEWARTII SUBSP. STEWARTII………………………………………….96
2.1 Introduction………………………………………………………………...96
2.2 Materials and methods…………………………………………………..99
xiv 2.2.1 Bacterial strains, plasmids, media and growth conditions……….99
2.2.2 HR and pathogenicity tests…………………………………………100
2.2.3 General recombinant DNA, genetic and bioinformatic techniques……………………………………………………………………101
2.2.5 Construction of Plac-hrpy, Plac-hrpS and Plac-hrpL plasmids……102
2.2.6 Construction of chromosomal P. stewartii hrp insertion mutants……………………………………………………………………….103
2.2.7 Site-directed mutagenesis of hrpY codon D57 by overlap extension
PCR and construction of missense mutant strains………………………104
2.2.9 Construction of P. stewartii strains with in-frame deletions in hrpX……………………………………………………………………….106
2.2.9 Construction of hrp-promoter reporter plasmids…………………105
2.2.10 Measurement of enzymatic reporter gene activity……………...107
2.2.11 Analysis of gfp reporter gene expression by flow cytometry….108
2.3 Results…………………………………………………………………..109
2.3.1 Sequence analysis of the hrp regulatory region of P. stewartii..109
2.3.2 Null mutant phenotypes and epistasis analysis…………………..112
2.3.3 Expression of hrp regulatory genes in different genetic backgrounds…………………………………………………………………114
2.3.4 Role of the conserved aspartate in HrpY………………………….117
2.3.5 Effects of environmental conditions on the expression of hrp genes in vitro……………………………………………………………………….118
xv 2.4 Discussion…………………………………………………………………120
2.5 Summary…………………………………………………………………..128
3. A NOVEL AUTOREGULATORY LOOP ACTIVATED BY HRPS
FROM THE HRPL PROMOTER MODULATES EXPRESSION OF THE
P. STEWARTII TYPE III SECRETION SYSTEM REGULATORS
HRPXY, HRPS AND HRPL...... 151
3.1 Introduction ...... 151
3.2 Materials and Methods ...... 156
3.2.1. Bacterial growth and media ...... 156
3.2.2 Molecular genetics and cloning techniques...... 156
3.2.3 Construction of polar hrpL interposon mutants and of plasmid
pMM191 ...... 157
3.2.4 Analysis of β-glucuronidase or β-galactosidase activity ...... 158
3.2.5 Total RNA isolation...... 158
3.2.6 Primer extension and DNA sequencing ...... 159
3.2.7 RT-PCR analysis ...... 159
3.2.8 Northern hybridizations...... 160
3.3 Results and discussion...... 161
3.3.1 Constitutive expression of HrpS indirectly upregulates hrpS
transcription by acting on the hrpL promoter ...... 161
3.3.2 The upstream region of the hrpL promoter contains a symmetrical
dyad required for HrpS-dependent activation ...... 163
xvi 3.3.3 Primer extension analysis defines the 5’ ends of hrpXY, hrpS and
hrpL transcripts and their potential promoters ...... 164
3.3.4 RT-PCR reveals that the hrpL-hrpXY and hrpXY-hrpS interoperon
regions are transcribed ...... 165
3.3.5 Northern blot analysis reveals the presence of long transcripts
spanning hrpL and hrpXY that are disrupted in a hrpL polar mutant .. 166
3.4 Conclusions...... 168
3.5 Summary...... 172
4. BIOCHEMICAL ANALYSIS OF THE RESPONSE REGULATOR
HRPY AND OF ITS ROLE AS A DUAL ACTIVATOR/REPRESSOR
OF HRP GENES...... 184
4.1 Introduction ...... 184
4.2 Materials and Methods ...... 187
4.2.1 Strains and growth conditions ...... 187
4.2.2 Construction of plasmid-borne gene fusions and of a hrpY[D57E]
allele for plasmid expression...... 188
4.2.3 Construction of plasmids for protein expression and
purification ...... 189
4.2.4 Purification of His6-HrpY proteins ...... 190
4.2.5 Mass spectrometry ...... 192
4.2.6 In vitro phosphorylation of HrpY ...... 193
4.2.7 Electrophoretic mobility shift assay ...... 194
xvii 4.2.8 DNase I footprinting ...... 195
4.2.9 β-glucuronidase enzyme assays ...... 197
4.2.10 Total RNA isolation ...... 197
4.2.11 Primer extension and DNA sequencing...... 198
4.2.12 Sequence analysis and modeling...... 198
4.3 Results...... 199
4.3.1 Deletion analysis of the regulatory elements activated or repressed
by HrpY...... 199
4.3.2 Overexpression and purification of HrpY proteins ...... 202
4.3.3 Phosphorylation of HrpY at D57 by Salmonella enterica
BarA198 ...... 204
4.3.4 Binding of HrpY to 5’ hrpS DNA fragments ...... 206
4.3.5 Binding of HrpY to 5’ hrpL DNA fragments...... 208
4.3.6 Effect of phosphorylation on in vitro HrpY binding activity .... 209
4.3.7 Effect of phosphorylation of HrpY on its in vivo activity ...... 210
4.3.8 Identification of the binding elements of HrpY by DNase I
footprinting ...... 211
4.4 Discussion...... 212
4.5 Summary...... 223
5. GENETIC ANALYSIS OF THE HRPX PAS-KINASE AND PARTIAL
CHARACTERIZATION OF RECOMBINANT HRPX PROTEINS AND
PROTEIN FRAGMENTS ...... 254
xviii 5.1 Introduction ...... 254
5.2 Materials and Methods ...... 255
5.2.1 Media and Strains and general microbial genetic techniques .... 257
5.2.2 Protein modeling ...... 258
5.2.3 Construction of strains with in-frame deletions in various
hrpX domains ...... 259
5.2.4 Construction of plasmids for genetic complementation tests .... 261
5.2.5. Construction of plasmids for expression and purification
of HrpX ...... 262
2+ 5.2.6 Overexpression and purification of His6 -HrpX protein by Ni -
NTA affinity chromatography ...... 264
5.2.7 Overexpression and purification of MBP-HrpX protein by
amylose affinity chromatography...... 266
5.2.8 Overexpression and purification of GST-PAS1-2HrpX protein
by glutathione affinity chromatography...... 268
5.2.9 Mass spectrometry protein fingerprinting...... 269
5.2.10 In vitro autophosphorylation of HrpX and
phosphotransfer reaction...... 270
5.2.11 UV-crosslinking of the HrpX input domain with [γ32P]ATP.... 270
5.2.12 Pathogenicity tests ...... 270
5.3 Results...... 271
5.3.1 The input region of P. stewartii HrpX is formed by two
PAS domains with low similarity between them ...... 271
xix 5.3.2 The two PAS domains of HrpX play different roles in
modulating its activity...... 274
5.3.3 HrpX does not transduce pH, osmolarity or many catabolic
metabolite-associated signals but it appears to sense stimuli related to
nitrogen metabolism and the Krebs cycle ...... 277
5.3.4 Expression and Purification of recombinant HrpX proteins ...... 279
5.3.5 Autophosphorylation kinetics and phosphotransfer ability of HrpX
...... 283
5.3.6 GST-PAS1-2 does not bind ATP...... 285
5.4 Discussion...... 286
5.5 Summary...... 293
6. N-ACYLHOMOSERINE LACTONE-DEPENDENT QUORUM-
SENSING IS REQUIRED FOR FULL EXPRESSION OF THE HRP-
TYPE III SECRETION SYSTEM OF PANTOEA STEWARTII SUBSP.
STEWARTII ...... 327
6.1 Introduction ...... 327
6.2 Materials and Methods ...... 332
6.2.1 Bacterial strains and plasmids...... 332
6.2.2 Genetic and molecular biology techniques...... 333
6.2.3 Plasmid construction...... 333
6.2.4 Enzyme assays...... 334
6.2.5 Plant assays ...... 334
6.3 Results...... 335 xx 6.3.1 An esaI mutant, containing a mutation in the AHL synthase gene,
shows a Hrp- phenotype...... 335
6.3.2 In epistasis experiments, constitutive expression of hrpS, but not
hrpY, suppresses the Hrp phenotype of esaI strains ...... 336
6.3.3 Expression of most hrp/wts genes is dramatically reduced in the N-
acyl-homoserine synthase mutant ...... 336
6.3.4 Expression of hrp secretion genes is population density-dependent
and EsaR mediates the effects of esaI ...... 337
6.4 Discussion...... 339
6.5 Summary...... 346
7. CONCLUSIONS……………………………………………………………………357
LIST OF REFERENCES……………………………………………………364
xxi
LIST OF TABLES
Table Page
1.1 Regulators of type III secretion systems...... 81
2.1 Bacterial strains and plasmids...... 130
2.2 Primers...... 134
2.3 Analysis of DNA and protein sequences ...... 136
2.4 Pathogenicity of wild type and hrp regulatory mutants and complementation analyse ...... 138
2.5 Effects of hrp regulatory genes on plasmid-borne hrp::uidA transcriptional fusions expressed in different genetic backgrounds …139
3.1 Bacterial strains and plasmids...... 173
3.2 Primers ...... 175
4.1 Bacterial strains and plasmids...... 225
4.2 Oligonucleotide primers...... 226
5.1 Bacterial strains and plasmids...... 295
5.2 Oligonucleotide primers...... 298
6.1 Bacterial strains and plasmids...... 348
6.2 Watersoaking symptom severity ratings and hypersensitive response elicitation caused by P. stewartii esaI-esaR mutants and epistatic effects of other Hrp regulatory genes...... 349
xxii
LIST OF FIGURES
Figure Page
1.1 Worldwide distribution of Stewart’s wilt……………………………….82
1.2 Stewart’s wilt disease cycle...... 83
1.3 The quorum sensing regulatory circuit controlling EPS synthesis in P.
stewartii...... 84
1.4 Bacterial secretion systems...... 85
1.5 Schematic representation of animal and plant pathogen type III
secretion apparatuses and of the flagellar basal body...... 86
1.6 Transcriptional organization of animal and plant type III secretion
systems..…………………………………………………………………………87
1.7 Regulatory circuits controlling type III secretion system transcription
in animal pathogens...... 88
1.8 Phytobacterial Hrp/type III secretion system regulatatory circuits. .. 89
1.9 Modular structure of two-component systems...... 92
1.10 Phosphorylation pathways in two-component systems...... 94
2.1 Physical map of the regulatory region of the P. stewartii DC283 hrp
gene cluster showing the hrpL, hrpX, hrpY and hrpS subclones used in this
study...... 140
xxiii 2.2 Analysis of the secondary structure of the IS element remnant located
in front of hrpS...... 142
2.3 PCR detection of the abortive IS element preceding hrpS in a
collection of P. stewartii strains ...... 143
2.4 β-galactosidase (β-Gal) and β-glucuronidase (GUS) activity of
chromosomal hrp-lacZ or hrpL-uidA fusions in different P. stewartii
genetic backgrounds, with and without various plasmid-borne Plac-hrp
regulatory genes...... 144
2.5 Expression of plasmid-borne hrp-uidA fusions in P. stewartii DC283.
...... 146
2.6 Flow-cytometry analysis of hrp-gfp fusion expression in P. stewartii
DC283 cells from infected corn plants compared to their expression in
LB...... 147
2.7 Effect of environmental conditions on the expression of various hrp-
lacZ fusions...... 149
2.8 Working model for the hrp regulatory cascade in P. stewartii...... 150
3.1. Physical map of the hrp regulatory region, mutants and plasmids. 176
3.2 Effects of HrpS+ constitutive expression on chromosomal hrp-lacZ fusions in various genetic backgrounds...... 177
3.3 The effect of deleting the upstream enhancer sequence (UAS) in the hrpL regulatory region on PhrpL expression...... 178
3.4 Primer extension analysis of hrpL, hrpX and hrpS transcripts...... 179
3.5. RT-PCR analysis of interoperon read-through transcription...... 180
xxiv 3.6 Northern blot analysis of hrp regulatory transcripts...... 181
3.7 Model of the hrp regulatory cascade including the autoregulatory loop triggered by HrpS...... 183
4.1 Expression of plasmid-borne hrpS gene fusions in P. stewartii...... 227
4.2 Expression of plasmid-borne hrpL gene fusions in E. coli SΦ200. . 228
4.3 Nucleotide sequence alignment of the hrpY-hrpS intergenic regions
from several erwinias and salient features of this region
in P. stewartii...... 229
4.4 Computer modeling of the intrinsic curvature of the hrpS promoter
region...... 231
4.5 Primer extension analysis and sequence analysis of the hrpL promoter.
...... 232
4.6 Overexpression and purification of HrpY...... 234
4.7 Mass Spectrometry analysis of recombinant HrpY
and HrpY[D57N]...... 236
4.8 Homology model of the crystal structure of P. stewartii HrpY ...... 237
32 32 4.9 Phospho-transfer of Pi from BarA198~ P to HrpY...... 239
4.10 Map of the hrpS regulatory region and the promoter fragments used as probes in gel shift experiments ...... 240
4.11 Electrophoretic mobility shift analysis of the binding of HrpY to the full length hrpS promoter...... 241
4.12 Electrophoretic mobility shift analysis of the binding of
HrpY to portions of the hrpS promoter...... 242
xxv 4.13 Competition analysis of the binding of HrpY to hrpS fragment C. 243
4.14 Map of the hrpL regulatory region and of the promoter fragments
used as probes in gel shift experiments...... 244
4.15 Electrophoretic mobility shift analysis of the binding of HrpY to
portions of the hrpL promoter...... 245
4.16 Electrophoretic mobility shift analysis of the binding of HrpY to the
full length hrpL promoter fragment...... 246
4.17 Effect of phosphorylation on HrpY binding to the hrpS and hrpL
promoters ...... 247
4.18 β-galactosidase activity of a chromosomal hrpS-lacZ fusion in a P.
stewartii hrpY null mutant carrying various plasmid-borne Plac-hrpY regulatory genes...... 248
4.19 DNase I footprinting assay of HrpY complexes at the
hrpS promoter...... 249
4.20 Model for hrpS activation...... 251
4.21. Model for hrpS activation...... 252
4.22 Model for hrpL regulation...... 253
5.1 Map of primers and PCR fragments used to construct hrpX mutations
(plasmids and chromosomal mutants)...... 299
5.2 Physical maps of the hrpX mutations and plasmids used for complementation studies...... 300
5.3 Plasmids used for expression and purification of HrpX...... 301
5.4 Plasmids used for expression and purification of HrpX domains.... 303
xxvi 5.5 Phylogenetic tree of HrpX homologs...... 305
5.6 PSI-PRED Secondary structure analysis of HrpX...... 306
5.7 Phylogenetic tree of PAS1-like domains in other two component
systems ...... 309
5.8 Threading of the PAS1 amino acid sequence into the FixL PAS
secondary structure...... 310
5.9 Structural model of the P. stewartii PAS1 threaded into the backbone
of B. japonicum FixL PAS...... 311
5.10 Complementation analysis and virulence phenotypes of P. stewartii
hrpX deletion mutants...... 312
5.11 The effect of various hrpX mutations on virulence and the expression
of hrpJ. a typical hrp secretion gene...... 314
5.12 The effect of environmental stimuli on hrpS-lacZ expression and
suppression of these effects by ectopic overexpression of
hrpY[D57N] ...... 315
5.13 Pilot experiment to overexpress polyHis-tagged HrpX proteins.... 317
5.14 Overexpression and purification of His6 -HrpX from E. coli
BL21(DE3) (pMM228)...... 318
5.15 Example of the results from nano LC-MS/MS protein fingerprinting of the tryptic fragment LHSSDTPIK from gel-excised His6 -HrpX...... 321
5.16 Overexpression and purification of MBP-HrpX by amylose affinity
and ion exchange chromatography...... 322
32 5.17 Kinetics of autophosphorylation of His6-HrpX by [γ P]ATP...... 324
xxvii 32 32 5.18 Transphosphorylation of HrpY by BarA~ P and His6-HrpX~ P.. 325
5.19 Purification of the GST-PAS1-2 protein from E. coli DH10B
(pMM374) and attempted direct photoaffinity labeling to [γ32P]ATP. .. 326
6.1 Maps of plasmid-borne reporter gene fusions used in this study. ... 350
6.2 Effect of an esaI mutation on the expression of several hrp regulatory and structural genes...... 351
6.3 Suppression of esaI regulatory effects on hrpJ by addition of
synthetic N-3-oxo-hexanoyl-homoserine lactone (OHHL)...... 352
6.4 Transcriptional expression of hrpJ as a function of cell density in
esaI, esaR and esaIR mutants...... 353
6.5 Effect of addition of synthetic N-3-oxo-hexanoyl-homoserine lactone
(OHHL) on expression of hrpJ...... 354
6.6 Model describing the RsmA regulatory network in P. carotovorum 355
6.7 Model for quorum sensing regulation of P. stewartii hrpS and
alignment of the putative esaR-like box (HrpS box)...... 356
7.1 Global regulation of hrp genes in P. stewartii…………………………..363
xxviii
LIST OF ABBREVIATIONS AND SYMBOLS
A: adenine
A: alanyl residue (Ala) aa: amino acid
A600: absorbance at 600 nm; see OD600
ATP: adenosine 5’-triphosphate avr: avirulence genes bp: base pair(s)
C: cytosine
CDD: conserved domain database
Ci: Curie (1 Ci=2.22x1012 dpm) cps: capsular polysaccharide synthesis genes cpm: counts per minute
CTD: carboxy-terminal domain
CUR: conserved upstream repeats
D: aspartyl residue (Asp)
DEPC: diethylpyrocarbonate
DPI: day post-inoculation dpm: disintegrations per minute dsp: disease specificity genes
xxix DR: direct repeats
DTT: dithiothreitol
E: E score, expected score
E: glutamyl residue (Glu)
ECF: extra-cytoplasmic function
EDTA: ethylendiaminotetracetic acid
EPS: extracellular polysaccharide esa: Erwinia stewartii autoinducer genes
F1: hybrid generated by crossing two parental lines
G: guanine
G: glycyl residue (Gly)
GC: guanine + cytosine content in molar %
GST: glutathion-S-transferase h: hour(s)
H: histidyl residue (His)
HEPES: 4-(2-hydroxy-ethyl)-1-piperazine-ethanesulfonic acid
HK: histidine kinase hop: Hrp outer protein gene
HR: hypersensitive response hrp: hypersensitive response and pathogenicity genes (pronounced
“harp”)
ICPPB: International Collection of Plant Pathogenic Bacteria
I.D.: columns internal diameter
xxx IHF: integration host factor
IMAC: immobilized metal affinity chromatography
IPTG: isopropyl-D-thiogalactopyranoside
IS: insertion sequence
K: lysyl residue (Lys) kb: 103 base pairs kcal: 103 calories
Kd : dissociation constant (in mol) kV: 103 volts kDa: 103 daltons or atomic mass units
M: in nucleic acid sequences, either A or C
M: in protein sequences, methionine (Met)
MALDI-TOF MS: mass absortion laser desorption ionization time of flight mass spectrometry
MBP: maltose binding protein
MES: 2-(N-morpholino)ethanesulfonic acid min: minutes mol: moles mRNA: messenger RNA
MS/MS: tandem mass spectrometry
MU: 4-methylumbelliferone
MUG: 4-methylumbelliferyl-glucuronide
MUGal: 4-methylumbelliferyl-β-D-galactopyranoside
xxxi m/z: mass over charge ratio
N: any base
N: asparagyl residue
Nano-LC: nanovolume liquid chromatography
NCBI: National Center for Biotechnology Information
NNPP: Neural Network Promoter Prediction nt: nucleotides
OD600: optical density at 600 nm; see A600
PAI: pathogenicity island
PAGE: polyacrylamaide gel electrophoresis
PAS: Per-Arnt-Sim
PCR: polymerase chain reaction
PFGE: pulsed field gel electrophoresis
PMSF: phenylmethylsulphonyl fluoride
QTL: quantitative trait loci/locus
Q-TOF: quadrupole time of glight (see MALDI)
RBS: ribosomal binding site rcs: regulation of capsule synthesis genes
RMSD: root mean square deviation
RR: response regulator rRNA: ribosomal RNA rsm: regulation of secondary metabolites
SD: standard deviation
xxxii SDS: sodium dodecyl sulfate
SK: sensor kinase
SE: standard error
T: thymine
T: in protein sequences, threonine (Thr)
TBE: Tris-borate EDTA buffer
TFA: trifluoroacetic acid
TMS: transmembrane sequence
Tris: tris(hydroxymethyl)aminomethane
tRNA: transfer RNA
TCS: two-component system
TTS: type III secretion
TTSS: type III secretion systems
UAS: upstream activating sequence
UTR: untranslated region
VUR: variable upstream region
W: either adenine or thymine
wts: watersoaking genes
X-Gal: 5-bromo-4-chloro-3-indolyl-β-D-galactoside
Y: in nucleic acid sequences, either cytosine or thymine
Y: in protein sequences, tyrosine (Tyr)
xxxiii
CHAPTER 1
LITERATURE REVIEW
1.1 STEWART’S WILT OF CORN AND ITS CAUSAL AGENT,
PANTOEA STEWARTII SUBSP. STEWARTII
1.1.1 Historical background
The history of bacterial wilt of sweet corn (synonyms: Stewart’s wilt of corn; Stewart’s disease of sweet corn and maize; bacterial leaf blight of maize; maize bacteriosis; Burrill’s disease of maize pro parte) is intimately linked to the dawn of bacteriology and phytopathology. The disease was first described by F. C. Stewart in 1897 in the bulletin of the
Agricultural Experiment Station, New York State, after he initially discovered the microorganism on infected corn plants in a Long Island
(New York) farm in 1895 (Stewart, 1897). A similar bacterial disease of corn was reported by T. J. Burrill eight years earlier (Burrill, 1889). This disease may have been Stewart’s wilt, however, Burrill, who was the first propose that bacteria could cause plant disease, failed to obtain a culture
1 of virulent microbes and his isolates were later identified by Theobald
Smith as nonpathogenic Bacillus cloacae (Smith, 1914). Stewart’s bulletin
is therefore considered to be the first report of the disease. In reality,
Stewart’s claims about the bacterial etiology of corn wilt were more
anecdotal than scientific, since they were merely based on the observation
that bacterial slime was constantly found in infected stems. Stewart was
indeed unable to reproduce symptoms on artificially inoculated yellow
dent corn, popcorn, teosinte and oats (Stewart, 1897) and satisfy what was
then called “Koch’s rules of proof”. His inconclusive experiments were
also plagued by contamination of control plants due to the use of infected
seed.
It was Erwin F. Smith who first proposed Pseudomonas stewartii nova
species as the causal agent of Stewart’s wilt in a paper at a meeting of the
American Association for the Advancement of Science on December 1898
(Smith, 1898). This report was followed by a more accurate description of
the cultural characteristics of this and other “uni-flagellate bacterial
pathogens of plants” (Smith, 1901). The formal proof of P. stewartii as
the causal agent was reported by E. F. Smith only in 1903 with an abstract
in Science (Smith, 1903) summarizing an address delivered in December
1902 before the Society for Plant Morphology and Physiology. For the
first time, Smith was able to artificially inoculate pure cultures of the
microbe on corn plants without wounding, via penetration of leaf stomata
and water-pores. He also conducted a detailed histopathological study,
2 defining the xylem of stems, leaves, husks and cobs as the primary
compartments colonized by the microbe, and parenchyma as secondary
site of infection. In his classic multi-volume treatise, “Bacteria in
Relation to Plant Diseases” (Smith, 1914), Smith summarizes much of the
work done on this microorganism during the first part of the last century.
Most of the literature following Smith’s pioneering work deals with the discovery of seed and insect transmission of the microbe (Smith, 1909;
Rand and Cash, 1924); attempts to control the disease using “lysins”
(later recognized as bacteriophages) (Thomas, 1938), insecticides (Ayers
et al., 1979; Sands et al., 1979), chemicals and antibiotics (Lockwood and
Williams, 1956; Natti, 1958; Pataky et al., 2000); prediction of disease
outbreaks based on weather variables (Stevens, 1934; Elliott and Poos,
1940; Castor et al., 1975); the use of disease resistant maize and sweet
corn lines (Pataky et al., 1998; Pataky et al., 2000); and the analysis of
the biochemical and quantitative genetic basis of the disease (Ming et al.,
1999). Initial genetic studies during the 1930-40s focused on the selection of virulent vs. avirulent strains with varying color and colony morphology after host passage (Wellhausen, 1937; Lincoln, 1939; Lincoln, 1947) and the analysis of mutation rates upon X-ray mutagenesis (Lincoln and
Gowen, 1942). As a historical curiosity, P. stewartii, along with
Drosophila and TMV, was used in one of the first studies aimed at defining the physical basis of genes and mutations (Gowen, 1941).
Systematic genetic studies of the microbe only began during the mid-
3 1970s with the isolation of temperature-induced, avirulent mutants
(Garibaldi and Gibbins, 1975) and the mobilization of F- and R-factors into P. stewartii (Gibbins et al., 1976; Coplin, 1978). During the 1980s, the introduction of bacteriophage Mu and Tn5 transposon mutagenesis allowed Coplin and associates to isolate capsular polysaccharide and water-soaking defective mutants (Coplin, 1979; McCammon et al., 1985;
Coplin et al., 1986). Such technical advancements were followed by an explosion of studies during the 1990s. The last 15 years have been characterized by the increasing use of genetic engineering and molecular biology to dissect cell biology, virulence and pathogenicity functions of this and several other model pathogens, to revise the taxonomic collocation of the microbe and to diagnose symptomatic and asymptomatic seed with added sensitivity and speed. This has gradually established P. stewartii, as not just an economically important pathogen, but as an extremely useful and powerful model for the analysis of basic gene regulation, cell-to-cell bacterial communication and virulence on corn, which is a monocotyledonous crop of enormous agricultural and industrial importance, with a vast community of researchers and a rich molecular genetics toolbox available.
1.1.2 Host range and resistance
Various species of the genus Zea are the common hosts of the disease, in particular sweet corn (Zea mays var. saccharata; the most susceptible
4 species), popcorn (Zea mays var. everta), and dent corn (Z. mays. var.
indentata; usually more resistant). The host range of P. stewartii includes
other monocotyledous plants. The pathogen has been isolated from
Eastern gama grass (Tripsacum dactyloides) and teosinte (Zea mexicana),
which is the Central-American progenitor of modern corn. Upon artificial
inoculation, P. stewartii is also able to produce symptoms or grow
endophytically in sorghum (Sorghum vulgare), Sudangrass (Sorghum bicolor), millet (Panicum miliatum), wheat (Triticum aestivum), oats
(Avena sativa) and other common grasses, such as yellow foxtail (Setaria glauca), black bent (Agrostis gigantea), Dactylis glomerata, Panicum capillare, Poa pratensis, Euchlena perennis, Setaria pumila, Tripsacum dactyloides, and T. zea. (Pepper, 1967; Ahmad et al., 2001). The only dicotyledonous plants experimentally infected are Golden Cluster beans
(Phaseolus vulgaris) and cucumber (Cucumis sativus) (Pepper, 1967).
Many of these plants may act as natural alternate hosts, contributing to the spread and over-wintering of the disease when sweet corn is not present (Poos, 1939).
Among the more than two thousand commercial dent and sweet corn hybrids, a wide range of disease responses exists, ranging from highly susceptible to highly resistant (Pataky et al., 2000). The use of resistant lines is the major control strategy for Stewart’s wilt. This resistance is
“partial” or “horizontal” (i.e. multigenic). Partial resistance is characterized by restriction of bacterial movement along the vascular
5 system (Braun, 1982). Initial studies identified two major genes and one
minor gene linked to resistance to Stewart’s wilt (Stevenson and Jones,
1953). Recently, one major QTL on chromosome S1 (locus sw1) and a
minor QTL on chromosome 9 controlling partial resistance have been
identified in the resistant line Ki14 (Ming, 1996; Ming et al., 1999). In
other lines, up to three QTLs on three different chromosomes could be
observed (Brown et al., 2001). Monogenic resistance to Stewart’s wilt
(also called “vertical”, or “race-specific” resistance) is not known to exist
in any corn accession (Stevenson and Jones, 1953; Pataky et al., 2000). In field trials, disease reactions of Z. mays inbred lines, F1 hybrids and sweet corn hybrids are classified by agronomists as resistant (R), moderately resistant (MR), moderately susceptible (MS), and susceptible
(S). In general, later maturing lines have higher levels of “horizontal”
(i.e. quantitative) resistance and, in these cases, the disease is confined to
the site of inoculation (in natural conditions, this is the feeding wounds
caused by the insect vector) and systemic wilt does not develop
(Suparyono and Pataky, 1989). Susceptible hybrids may show systemic infection when artificially inoculated as late as the 7- to 9-leaf stages, but
systemic infection of resistant hybrids is not common after the 3-leaf
stage (Pataky et al., 2000). Current national estimates of yearly losses from Stewart’s wilt are not available. In the past century, disease losses
increased progressively until 1930-1932, when the most spectacular
epidemics were recorded. Field losses on sweet corn and popcorn were
6 about 13%. During subsequent years, the increasing use of resistant
hybrids and varieties reduced the losses to a trace, except for occasional
outbreaks (Pepper, 1967). This pattern has continued, except when
individual sweet corn growers plant susceptible, early maturing sweet
corn varieties for marketing reasons. Following mild winters, heavy beetle
populations may overwinter in sheltered fields and these growers still
suffer devastating losses. Control of beetle populations by insecticide sprays or seeds treatments is the second major strategy of disease control currently available (Heichel et al., 1977; Ayers et al., 1979; Munkvold et al., 1996; Pataky et al., 2000; Kuhar et al., 2002). Yield losses can be
significant in susceptible varieties and in early infections because of the
high incidence of systemic infection in such cases. When plants are
infected systemically, yield is reduced about 1% for each 1% increase in
incidence of systemically infected plants, and this usually translates into a
complete destruction of the individual plant ears. In disease-conducive
years, field incidences of 40-100% can be observed (Pataky, 2003).
1.1.3 Geographic distribution of the disease
Stewart’s wilt probably originated in North America, where it was first
described. Moreover, it was not described (Pepper, 1967) in the centers of
origin of corn (Central America and Mexico) until the mid 1900s.
Considering the ability of P. stewartii to live as an endophyte in grasses
and the wide host range of the corn flea beetle, one may speculate that the
7 introduction of sweet corn into North America by native Americans
provided the occasion for a host species jump, analogous to what is
thought to have happened for Erwinia amylovora (Van der Zwett and Keil,
1979) upon the introduction of cultivated Pomaceae by the first American
settlers during the 1600s. In this respect, and interestingly from an
evolutionary point of view, it is notable that another erwinia, P.
agglomerans pv. gypsophiliae (Cooksey, 1986), also originated in the US
and that all three species share a highly conserved pathogenicity island
located on their chromosome or on a plasmid.
The occurrence of Stewart’s wilt epidemics appears to be strictly linked
to the presence of the most common insect vector, Chaetocnema pulicaria
(Pepper, 1967), or the related species C. denticulata and Diabrotica
undecimpunctata, which are present only in the mid-Atlantic to the
eastern portion of Mid-Western USA. The absence of these vectors is
commonly believed to be the reason that the disease has not spread to
other areas, even though it can be seed-borne. In the USA, the disease is endemic from the Mid-Atlantic states, through the Ohio Valley, to
Missouri and Iowa, with infrequent outbreaks 500 km north and south of the “Corn Belt” (Pataky 2000). This area includes Arkansas, Delaware,
Illinois, Indiana, Kentucky, Maryland, Missouri, New Jersey, New York,
Ohio, Pennsylvania, Tennessee, West Virginia and Virginia. Outbreaks
were also reported in Idaho and Washington state in 1920, but the disease
did not become established, possibly because of the dry climate and/or the
8 absence of the insect vector (Pepper, 1967). From the USA, the pathogen has been spread worldwide during the last 100 years, probably via trading of contaminated corn seed, mainly causing isolated field outbreaks or quarantine interceptions of infected material in Central America, Europe, and Asia (Fig. 1.1). The first case of disease outside of North America occurred in Italy (Pasinetti, 1937), with recurrent outbreaks in 1937,
1948, 1982 and 1983 (Anonymous, 1983), which were all linked to US- imported seed. Based on the European Plant Protection Organization
(EPPO) Data Sheets (EPPO, 1999), the disease has also been reported in
Viet Nam, Thailand, India, China (1942), Malaysia, Austria, Greece,
Poland (1942), Romania, Russia (1932), Switzerland (1950), Canada
(1931), Costa Rica (1949), Puerto Rico (1927), Mexico (1938, 1996,
1999), Guyana, Brazil (1968), Peru and Bolivia, but in most cases
Stewart’s wilt has not become endemic in the new environments. Notable exceptions are the sporadic outbreaks on sweet corn reported in Italy (Po
Valley; C. Bazzi, pers. commun.), Austria (in the regions of
Niederosterreich, Burgenland and Steiermark) and Mexico (Toluca Valley,
Oaxaca, Tabasco, Tlaxcala, Vera Cruz). In these countries, it is not known how the disease spread from the initial foci and became established, but such cases perhaps show a potential ability of P. stewartii to adapt to new secondary host-vector combinations (Pataky, 2000). Based on international agreements, countries with reported outbreaks of bacterial wilt must undergo strict field inspections for seed destined to export
9 markets. The seed from infected fields must be tested prior to export by a
certified lab and certified as pathogen-free. This requirement has
generated much research on diagnostics applied to seed lots. The
regulation of the corn seed trade and the P. stewartii quarantine rules
(European Council Directive 2000/29/EC, May 8, 2000) are major issues
in the turbulent Europe-USA agricultural trade relationships.
1.1.4 Symptoms and signs
Plants may be infected at any developmental stage and the bacterium
can cause a wide range of symptoms, which may be confused with
drought, potassium or nitrogen deficiency, insect damage, or fungal
diseases (e.g. Northern corn leaf blight, Southern corn leaf blight,
Helminthosporium leaf spot). In addition, three other bacteria cause
diseases on corn, including Pseudomonas alboprecipitans (bacterial leaf
blight), P. syringae pv. syringae (holcus spot) and P. andropogonis
(bacterial stripe), but their distribution and importance is extremely
limited. It would be interesting to determine whether or not common virulence factors control the development of water-soaking symptoms in these other gram negative corn pathogens.
Stewart’s wilt typically occurs in two phases. First, infected seedlings
or young plants show leaf streaks and water-soaked lesions with wavy,
irregular margins, from 1 to 10 mm wide and as long as the leaf. These
lesions quickly become necrotic, leading to leaf blight and eventually
10 wilting (Fig. 1.2). Under warm and humid conditions, bacterial ooze may appear on the surface of the lesions. Such early infections can be very severe and often cause complete destruction of the plants. Plants that survive early infections are stunted and later may form premature, abnormal ears, with tassels that are dwarfed, whitened and dead.
In the second phase of the disease, which typically occurs in mature plants that are infected after tasseling, the infected leaves show characteristic streaks parallel to the main veins (Fig. 1.2). Late infections are more common in susceptible dent corn lines. The lesions again have wavy irregular margins and are from yellow to light green in color. With time, the lesions turn brown and coalesce, causing leaf blight. Typically, insect feeding scars are associated with the lesions and they indicate the entry point of the pathogen. The size of such lesions is considerably smaller in resistant varieties, where they often appear as oval spots, 1-2 inches long (Pepper, 1967). Discolored, rotten cavities full of bacterial slime can be observed in the pith near the crown of mature stalks.
Similarly, light microscopy of transverse sections of infected stalks or leaf lesions may reveal yellow bacterial ooze extruding from the vascular bundles. This is a good diagnostic test in field examinations. Severe
Stewart’s wilt infection may predispose plants to stalk rot disease
(Pepper, 1967).
11
1.1.5 Source of inoculum, disease cycle and histopathology
The pathogen can overwinter in the intestine of a few insects of the order Coleopter.. Infected seeds are the other major means of overwintering (Smith, 1909). The importance of secondary hosts in the disease cycle is not known, but given the ability of P. stewartii to grow endophytically in many grasses (Ahmad et al, 2001), we cannot dismiss this possibility. Soil and infested crop residues are not relevant to pathogen survival and overwintering (Pepper, 1967).
The insect vectors belong to the order Coleoptera, family
Chrysomelidae. The most important species are the corn flea beetle (C. pulicaria), the toothed flea beetle (C. denticulata) and the 12-spotted cucumber beetle (D. undecimpunctata). The first experiments pointing to insects as the most important factor in Stewart’s wilt epidemiology date back to the 1920s (Pepper, 1967) and this relationship was proved conclusively by Elliott and Poos (Elliott and Poos, 1934). Extensive experimentation during the 1930s. Thousands of different species were tested as vectors for P. stewartii, other insect species were identified as potential carriers of the disease, including the Northern corn rootworm
(D. logicornis), Western corn rootworm (D. virgifera), the larval stage of
May beetles (Phyllophage sp.), seed corn maggot larvae (Hylemya cilicrura), and the common wheat wireworm (Agriotes mancus) (Elliott and Poos, 1940). However, under field conditions, C. pulicaria appears to
12 be the prevalent vector of the disease (Elliott and Poos, 1940).
Interestingly, C. pulicaria, C. denticulata and D. undecimpunctata have never been reported outside North America, which may explain the limited distribution of the disease outside of the US.
Infected seeds are another known mean of overwintering. A comprehensive review of the literature on seed infection is found in a recent report by Pataky (2000). In corn kernels, bacteria colonize the chalazal region, the aleurone layers and the apoplastic spaces of the endosperm, but are not found in the embryos or seed coat (Rand and Cash,
1924; Ivanoff, 1933). Survival in the seed varies from 5 to 8 months after harvest, depending on the storage temperature (Rand and Cash, 1924).
From infected seed, the bacteria can enter the seedling, with various degree of efficiency depending on the severity of infection in the parent plant and the susceptibility of the host to systemic infection (Pataky,
2000). The early literature mentions plant-to-seed infection rates (kernel infection rate) ranging from 2 to 85% (Pepper, 1967). However, more recent studies have shown that the rate of kernel infection is below
0.025% when the parent is resistant, and ranges from 0.25% to 10% when the parent is rated as susceptible or highly susceptible; the highest rates are limited to cases where the parent plant is systemically infected
(Pataky, 2000).
The disease cycle of Stewart’s wilt is depicted in Fig. 1.2, which summarizes our current understanding of P. stewartii
13 biology/epidemiology. The primary source of inoculum is bacteria over-
wintering in adult corn flea beetles (Fig. 1.2, step 2) or, much less
frequently, infections are started by bacteria surviving in the storage
tissues of maize kernels (Fig. 1.2, step 1). In the first case, the incidence of field infection is correlated with the size of the surviving population of insects. In this respect, studies conducted early last century (Stevens,
1934), based on 35 years of data, showed that Stewart’s wilt infections do not occur when the sum of the monthly mean temperatures for December through February is below 90°F, but disease incidence is high when the sum exceeds 100°F, due to higher populations of surviving vectors.
Penetration of healthy host tissues occurs externally via insect feeding wounds (Fig. 1.2, step 4) or endophytically from the germinating infected seed (Fig. 1.2, step 3). Under experimental conditions, the pathogen can also enter through stomata and hydatodes (Pepper, 1967). It is not clear whether bacteria are transmitted from the insect to the plant via an oral or a fecal route, but bacteria are probably associated with the alimentary tract. In general, very little is known about P. stewartii-insect vector
relationships. With the vast array of molecular tools now available, this
could represent an area of active research in the future. Following
wounding and inoculation, the bacteria localize in the intercellular spaces
of the parenchyma and in the embolic xylem vessels, probably starting
multiplication on the few nutrients available in the cell wall fluids (Fig.
1.2, step 6, 9). In this respect it is worth remembering that like most
14 bacterial plant pathogens, P. stewartii remains in the apoplast and never enters living host cells. This is in contrast to many mammalian pathogens that invade host cells. Based on studies by Braun (Braun, 1982), bacteria inoculated by vacuum infiltration remain singular or in small groups from
1 to 3 h after entering the tissue. Very little EPS seems to be produced during this early phase. A well-formed capsule surrounded by abundant slime could be seen at 12 h (Braun, 1982). Eventually, active multiplication results in the formation of microcolonies in the intercellular spaces of the parenchyma, which can be seen at 30 h post- inoculation. Appearance of watersoaked lesions and blight symptoms
(possibly a form of plant cell death/cytotoxicity) are visible from 48 h to
5 days post-inoculation. Bacteria colonize also the xylem vessels.
Whether bacteria actively force openings into vessels via enzymatic activity or passively break vessel pits by multiplication and mass swelling is not known (Braun, 1982). Production of pectolytic enzymes, which would be required for active movement across the xylem pits, is not observed in vitro (Braun, 1990), but it is not known whether they are produced in vivo. In experiments using 32P-labelled cells, it was shown that the rate of colonization of the xylem was correlated with the rate of transpiration and that the extent of the colonization did not differ between resistant and susceptible plants (Warren, 1951). This result implied that, at least under those experimental conditions, the free mass-flow of bacteria in the xylem was due to physical movement rather than
15 multiplication. Systemic colonization of the vessels causes wilt
symptoms, which is more dramatic in highly susceptible sweet corn
varieties. In susceptible plants, rapid systemic infection, plugging of the
xylem by EPS and fast deterioration and invasion of the adjacent
parenchyma cells occurs. The final result is often the destruction of the
whole vascular bundle and the appearance of wilt symptoms (Wellhausen,
1936; Braun, 1982; Braun, 1990). Bacteria may exit into the intercellular
spaces of the mesophyll through the primary cell wall of broken xylem
vessel pits during the late stages of pathogenesis (Braun, 1982). Work
done in the early 1980s has shown that specific corn agglutinins
(Bradshaw-Rouse et al., 1981) and higher concentrations of methoxy-
benzoxazolinone (Whitney and Mortimore, 1961) may be involved in
restricting massive colonization of parenchyma and xylem in resistant
lines.
1.1.6 Taxonomy
P. stewartii belongs to the domain Bacteria, phylum B.XII
“Proteobacteria phy. nov.”, class III “Gamma-proteobacteria”, order XII
“Enterobacteriales”, family I “Enterobacteriaceae”, genus XXIII
“Pantoea” (based on the 2000 Bergey’s Manual Trust, www.cme.msu.edu/bergeys). The nomenclature of this bacterium has
changed over time as a consequence of evolving methods of bacterial typing and different philosophical approaches to bacterial taxonomy. The
16 current proposed name is Pantoea stewartii (Smith 1898) subsp. stewartii
Mergaert, Verdonck & Kersters 1993, comb. nov., basonym Erwinia stewartii Smith 1898, Dye 1963 (Smith, 1898; Dye, 1963; Mergaert et al.,
1993). The basonym is still considered valid because it was included in the latest “Approved Lists of Bacterial Names” (Skerman et al., 1980).
The type strain defining the species is ATCC 8199 (Holt, 1984), deposited at the American Type Culture Collection as ICPB SS11 (originally classified as Xanthomonas stewartii Smith Dowson and isolated in the
USA from sweet corn by W.H. Burkholder in the 1940s). Other synonyms found in the literature over the last 105 years are "Pseudomonas stewartii" Smith 1898, "Bacterium stewartii" (Smith 1898) Smith 1911,
"Aplanobacter stewartii" (Smith 1898) McCulloch 1918, “Bacillus stewartii” (Smith 1898) Holland 1920, "Phytomonas stewartii" (Smith
1898) Bergey et al. 1923, "Xanthomonas stewartii" (Smith 1898) Dowson
1939, "Pseudobacterium stewartii" (Smith 1898) Krasil’nikov 1949
(Source: List of Bacterial Names with Standing in Nomenclature: Société de Bactériologie Systématique et Vétérinaire, http://www.bacterio.cict.fr/).
In 1898, Erwin F. Smith provided the first description of the cultural characteristics of the bacterium. Originally it was erroneously described as motile with a single polar flagellum and was therefore placed in the genus Pseudomonas (Smith, 1898; Smith, 1901). In 1914, E. F. Smith reclassified the species as Bacterium stewartii (Smith, 1914). After
17 McCulloch reported that it was nonmotile and lacked flagella in 1918, it
was reclassified as a member of the genus Aplanobacter (McCulloch,
1918). An attempt by Holland to compile updated and comprehensive generic indexes of bacteria resulted in Aplanobacter stewartii being moved into the genus Bacillus (Holland, 1920). This lasted until the first edition of Bergey’s Manual was published in 1923 and this bacterium was then included in the genus Phytomonas, which encompassed all gram- negative plant pathogens (Bergey and al., 1923). In 1937, the use of the genus name Phytomonas was questioned by Elliott because it was already in use by zoologists (Elliott, 1937). Then in 1939, Dowson proposed that
P. stewartii be placed in the genus Xanthomonas based on its biochemical properties and the presence of a yellow pigment (Dowson, 1939). This proposal ignored the facts that P. stewartii was nonmotile, did not produce the same yellow carotenoid and was facultatively anaerobic. In
1948 in the 6th edition of Bergey’s Manual, Burkholder classified P.
stewartii as a member of the genus Bacterium, a quite artificial and
heterogeneous taxon (Breed and al., 1948). Only in the 7th edition of
Bergey’s Manual, published in 1957, did Burkholder accept the
nomenclature Xanthomonas stewartii, but under the postil species incerta sedis (Burkholder, 1957). Between 1962 and 1963, the New Zealand scientist Douglas Dye revisited Xanthomonas stewartii taxonomy, accepting early suggestions that the microbe was related to coliform bacteria (Waldee, 1941) and a degenerate member of the
18 Enterobacteriaceae family. Consequently Xanthomonas stewartii was
classified in the genus Erwinia, as member of the so called “herbicola” group III, along with other “yellow” erwinias such as Erwinia herbicola
(syn. Enterobacter agglomerans; comb. nov. Pantoea agglomerans),
Erwinia ananas and Erwinia citri, and apart from the “amylovora” group I
and “carotovora” erwinia group II (Dye, 1963). Dye’s nomenclature is
accepted by the latest “Approved Lists of Bacteria Names” (Skerman et
al., 1980), and is used in the IXth edition of Bergey’s Manual (1984).
More recently, however, this official classification was challenged by
Mergaert et al. (1993). They proposed to move the group III erwinias into
the genus Pantoea based on DNA hybridization data and numeric
taxonomy. Consequently they renamed E. stewartii as Pantoea stewartii
subsp. stewartii. Subsequent analyses of the 16S rRNA gene sequences of
16 type species of the genus Erwinia (Kwon et al., 1997) confirmed the
distribution of most Erwinia species into four clusters, fundamentally
corresponding to Dye’s groups (with the exception of Erwinia salicis,
Erwinia rubrifaciens, and Erwinia nigrifluens, which form a fourth and distantly related clade). E. stewartii grouped with other erwinias of Dye’s
“herbicola” group in cluster I, even though the bootstrap value between it and the “amylovora” branch was only moderate. In the same study, the authors found that other enterobacteria, such as Escherichia coli,
Klebsiella pneumoniae, and Serratia marcescens, were intermixed with species of Erwinia. If nothing else, this finding raises questions on the
19 definition of bacterial species in this family and on the sole use of 16S rDNA analysis to reconstruct phylogenetic relationships. Another 16S rDNA study by Hauben et al. (1998) used 29 enterobacterial plant pathogens and likewise supported the splitting of Dye’s genus Erwinia into four groups, which they proposed to make different genera: Erwinia
(Dye’s group I), Pectobacterium (Dye’s group II), Pantoea (Dye’s group
III), and Brenneria (Kwon’s group 4). They also confirmed the collocation of the Stewart’s wilt agent into the monophyletic genus
Pantoea (Hauben et al., 1998).
P. stewartii is a yellow gram-negative rod, 0.9-2.2 µm in length by 0.4-
0.8 µm in diameter, that is nonmotile and non flagellate (Pepper, 1967;
Kado and Coplin, 2001). In this regard, P. stewartii has been recently reported to be capable of swarming in soft agar motility plates but the cellular basis for this phenotype is not known (S. von Bodman, pers. commun.). P. stewartii is facultatively anaerobic, cytochrome c oxidase negative, nitrate reductase negative, catalase positive and urease negative.
It can assimilate organic and inorganic nitrogen (ammonium ions and nitrates). The bacterium is able to produce acids from galactose, mannose, sucrose, glucose, mannitol, arabinose, xylose, fructose, glycerol, inositol and sometimes from sorbitol, whereas it produces alkali from rhamnose and dulcitol (Pepper, 1967; Kado and Coplin, 2001). Strains of P. stewartii were reported by Dye as delayed lactose fermenters (Dye, 1963), but this phenotype is not usually present (Kado and Coplin, 2001). These
20 differences could reflect strain divergences. P. stewartii shows a weak
endogenous and uncharacterized ß-galactosidase activity, easily visible in
rich media containing X-Gal, but not in McConkey-lactose media. Given that X-Gal is known to be non-specifically hydrolyzed by several
hydrolases (Neidhardt, 1993), such activity does not necessarily imply the
existence of a ß-galactosidase (lacZ) gene. The bacterium is also unable
to hydrolyze starch, casein, aesculin, lipids and pectins, but it can weakly
liquefy gelatin in a strain specific manner (Dye, 1963). It utilizes
gluconate and tartrate, but not propionate, and it can use asparagine as a
sole source of C and N (Dye, 1963). The bacteria are salt tolerant and can
grow in media with up to 7% NaCl (Burkholder and Starr, 1948). The
thermal death point is 53°C, maximum growth temperature is 39°C,
optimum temperature is 28-30°C, and minimum growth temperature is 8°C
(Dye, 1963). P. stewartii is able to survive for 14 months in agar stabs at
15°C but for only 7-10 days on agar plates at 4°C (Pepper, 1967).
1.1.7 Molecular Diagnostics
A comprehensive review of the methods used for the detection and
typing of P. stewartii is available in a recent monograph on phytobacterial
diagnostics (Kado and Coplin, 2001). Common methodologies for
detection and/or identification of P. stewartii involve simple field
inspection of seed plants, grow-out tests of seed, and the use of selective
and semiselective media to isolate pure cultures of bacteria. Cultures are
21 subsequently identified by classic morphological and physiological tests,
analysis of membrane fatty acid methyl-ester derivatives by gas
chromatography (GC-FAME; Wells et al., 1994), and/or enzyme-linked
immunosorbant assays (Lamka et al., 1991). In the last ten years, several
DNA based methods have been developed. One of the first reports described the use of arbitrary primer PCR to produce a species-specific
DNA probe (Blakemore et al., 1992). This approach was subsequently expanded by developing nested primers that amplified the same DNA
fragment (Blakemore et al., 1999). In 1994, a report on the use of the ligase chain reaction and 16S rRNA primers to identify P. stewartii was
published (Wilson et al., 1994), but the use of radioisotopes limited such
methods for routine assays. More recently several other PCR tests have
been reported for the purpose of genetic diversity studies (Fessehaie et al., 2002; Waleron et al., 2002) or detection/identification (Coplin et al.,
2002). In general, P. stewartii strains show a greater homogeneity in their genetic fingerprints than other pantoeas. This is analogous to the situation with E. amylovora, another species that originated from North America.
Genomic analysis of XbaI or SpeI macrorestriction fragments using pulsed field gel electrophoresis showed that P. stewartii strains are highly homogeneous, with Nei and Li similarity coefficients ranging from 0.86 to
0.9 (unpublished; calculated from Coplin et al., 2002). This technique could be used to differentiate P. stewartii from P. agglomerans, P. ananas and Pectobacterium carotovorum.
22
1.1.8 Genetics
The P. stewartii genome has a mol%G+C content of 54.6-55.1%
(estimated for two strains only; Holt, 1984). The size of the P. stewartii
chromosome is estimated to be between 3.5 and 4 Mb by PFGE analysis of
genomic macrorestriction fragments (Coplin et al., 2002). This estimate
does not include most of the 8 to 13 resident plasmids with sizes ranging
from 4 to 318 kb that are present in each of at least 31 virulent strains
from a geographically representative collection (Coplin et al., 1981).
Plasmid DNA may account for up to about 880 kb of the genome
(calculated from data reported in (Coplin et al., 1981). Eight class sizes
(4, 13, 25, 44, 65, 74, 106, and 318 kb) were common among 87% of more of the strains. Campbell-type recombination among replicons could account for “missing” and new plasmids in any given profile (Coplin et al., 1981). The presence of the 318 kb plasmid is somewhat in doubt because the newer plasmid purification techniques presently used in our lab do not isolate it. However, it is of particular interest to us, because several lines of experimentation suggest that the hrp/wts PAI may be located on it. This would be similar to what has been found for the hrp
cluster in P. agglomerans pv. gypsophilae, which is carried on a 150 kb pPATH plasmid (Manulis et al., 1991; Nizan et al., 1997).
In avirulent laboratory strains that have been in culture for over 40 years, the number of plasmids was reduced, but still most of the eight
23 conserved classes were present (Coplin et al., 1981). Several plasmids from strain SW2 have been totally or partially characterized. Plasmid pSW100, a 4.2-kb low copy number (2 copies per cell) mobilizable replicon, has been completely sequenced and found to contain a 0.7 kb origin of replication that was homologous to orgins from p15A, ColE1 and
ColA (Fu et al., 1995). Plasmid pSW500 (a 35-kb low copy plasmid) contains seven 16-bp iteron repeats, a DnaA box and a 1005-bp repA gene
(Fu et al., 1996). The replication origin of pSW800 has been characterized as well (Wu et al., 2001) and the results suggest that a mechanism similar to that regulating plasmid replication in the IncB, IncIa and IncL/M groups also regulates pSW800 replication. Plasmid pSW1200 (106 kb) has an approximate copy number of one and contains a bacteriophage P1-like ori (with a repA gene and eight 19-bp iteron sequences). Plasmid pSW1200 belongs to the IncY group (Fu et al., 1997). The significance of the coexistence of so many replicons in the same cell is both puzzling and intriguing. Perhaps they play a role in adaptation to various niches (such as secondary hosts and insect vectors) (Coplin et al., 1981). No clear
association of any P. stewartii plasmid with virulence or specific
phenotypes is known. The genome of P. stewartii strain DC283 (a
spontaneous NalR derived by D. L. Coplin from the virulent strain ICPPB
SS104 isolated by Art Hooker in 1967) is currently being sequenced at
Baylor College of Medicine and Wisconsin Genetic Center using a random
shot-gun library approach.
24 The genetic techniques available for P. stewartii are quite similar to
that of other enterics. P. stewartii can exchange plasmids with E. coli; it
supports replication of phage Mu (but is not infected by Mu virions) and
of many plasmid replicons, such as ColE1, p15, IncP1, and IncW groups.
However, F-factors are unstable and exhibit fertility inhibition, but not
surface exclusion, suggesting they are probably incompatible with one of
the indigenous plasmids (Coplin, 1978). Lysogenic phages are present in
about half the strains tested (D. L. Coplin, unpublished), but they have
never been genetically characterized. P. stewartii is insensitive to bacteriophage lambda infections, probably due to the absence of the LamB receptor, and infections by phage P1 or P22, two common phages used in generalized transduction of enterics. Although transduction systems are not available for genetic studies of P. stewartii, transposon mutagenesis has been an important tool in dissecting P. stewartii virulence. Delivery of transposons into P. stewartii cells was initially quite difficult because many of the tools available for other enteric bacteria would not work properly in this species. Widely used broad host range, suicide delivery vectors, such as pJB4JI, carrying modified Mu or Tn5, contrary to what observed in P. agglomerans subsp. herbicola, Pectobacterium chrysanthemi and P. carotovorum, replicated well in P. stewartii (Coplin,
1979). Temperature sensitive replicons, such as pUT13::Tn5 (used successfully in P. chrysanthemi) could not be used in P. stewartii because it does not grow at the non-permissive temperatures needed. Direct
25 infection with mini-Mu or MuS::Tn9 was not feasible because P. stewartii
is not sensitive to Mu(G-) phages. One of the first successful strategies for insertion mutagenesis was the use of an RK2::Mu cts62 hybrid plasmid
(pRK210). The replication and transposition of this Mu derivative was
inducible at 42°C after conjugal transfer from E. coli. (Coplin,1979). This
strategy was used to successfully isolate a number of of auxotrophic
mutants (Coplin, 1979). A second approach made use of a hybrid plasmid
comprised of the cryptic P. stewartii conjugative plasmid pDC250
containing a Tn10 insertion (pDC251) and phage Mu kan pf7701. This
genetic strategy allowed the isolation of several avirulent insertion
mutants affected in exopolysaccharide (EPS) synthesis and water-soaking
(Wts) ability for the first time (McCammon, 1985). Later, random
mutagenesis and the isolation of new Wts- mutants was achieved by using pMO75, an IncP-10 suicide replicon from Pseudomonas aeruginosa, to
deliver a Tn5 transposon (Coplin et al., 1992). The current standard
procedure for random mutagenesis in P. stewartii makes use of mob+, R6K
ori, pir-dependent suicide replicons in pUT vectors, i.e. the mini-Tn5
series of transposons for polar insertions and gene/protein fusion
generation (de Lorenzo et al., 1990; Majerczak, Merighi and Coplin,
unpublished) or derivatives thereof. The transposons are delivered via
biparental matings from E. coli SM10 or S-17 λpir. Genomic libraries
were constructed in E. coli using broad host range, mobilizable cosmid
vectors, such as pVK100 and pLAFR3 (Coplin et al., 1986; Dolph et al.,
26 1988). These vectors are relatively large (20-22 kb), which limits the
average insert size to 25-30 kb. Following mutagenesis of the cosmids, or subclones thereof, allele exchange is performed by well-established genetic techniques for gram negative bacteria (Maloy et al., 1996). One common approach that we follow when using mutagenized clones with large inserts with at least a few kb of flanking homologous sequences is selectable marker exchange by plasmid incompatibility (Ruvkun and
Ausubel, 1981). The clone, usually a pLAFR derivative (Friedman et al.,
1982), is mobilized by biparental (Simon et al., 1983) or triparental (cit; using pRK2013 or pRK2013::Tn7 as helper plasmids) matings (Ditta et
al., 1980) into P. stewartii. Subsequent destabilization of the plasmid and
forced single cross-over with the homologous region of the chromosome is
done by mobilizing and selecting also for a second incompatible plasmid
(e.g. pJP4JI GmR or pR741 TpR) (Ruvkun and Ausubel, 1981). More recently, we introduced the use of R6K suicide plasmids followed by positive selection in presence of fusaric acid to loose the vector encoded tetracycline resistance (pLD55; Metcalf et al., 1996). This has allowed the construction of unmarked (substitutions and deletions) or marked (Ω or aphA3 insertions) mutations in P. stewartii. As little as 600 bp of flanking sequence homology is needed to exchange the allele into the P. stewartii chromosome (Merighi et al., 2003), obviating the problematic use of sacB-based positive selection vectors, which work poorly in P. stewartii.
27 1.1.9 Molecular pathogenesis
P. stewartii is a vascular pathogen that can also grow in the
intercellular spaces of the parenchymatic tissues of leaves. A major mechanism of virulence (in sensu lato) is represented by the production of exopolysaccharide (EPS) slime, which occludes xylem vessels causing the plant to wilt (Ivanoff et al., 1938; Braun, 1982). EPS is required for biofilm formation and rapid systemic movement in the plant: EPS- deficient strains grow and spread more slowly in the host vascular tissues, are less likely than the wild-type to colonize plants systemically and are unable to cause wilting in infected plants (Braun, 1990). The EPS of P.
stewartii is a very large (~45 MDa), viscous, highly charged
heteropolysaccharide composed of glucose, galactose and glucuronic acid in a 4:2:1 ratio. No other EPSs are made in culture. The EPS forms both a
bound capsule and loose slime. Capsule synthesis is constitutive but
additional slime production is induced by the availability of free sugars in
the growth medium. In addition to its involvement in virulence, EPS
possibly plays a role in protection from phytoagglutinins in the vessels
(Bradshaw-Rouse et al., 1981) and in the colonization of its insect vector.
The genes involved in capsular polysaccharide synthesis were initially cloned in cosmid pES2144 based on complementation of spontaneous non- mucoid mutants (Coplin et al., 1986). Cosmid pES2144 contains five complementation groups (cpsA-E) and the galE gene. Most of these genes are conserved in E. amylovora and E. coli based on sequence homology
28 (Bernhard et al., 1993) and interspecific complementation tests (Bernhard
et al., 1996), except for several glycosyl transferase genes that may account for differences in the structure the two polysaccharides. Cloned
E. amylovora ams genes complemented most P. stewartii cps mutants for slime production and virulence in corn seedling (Bernhard et al., 1996),
but the arrangement of the genes in the two clusters is slightly different.
The P. stewartii cps gene cluster is structurally and functionally related to
the group I capsule biosynthetic gene clusters of other enteric bacteria
(e.g. the colanic acid cps cluster in E. coli) (Dolph and Coplin, 1987;
Dolph et al., 1988). Regulation of the P. stewartii cps genes also appears
to be similar to that for colanic acid in E. coli (Torres-Cabassa et al.,
1987) and amylovoran in E. amylovora. The Rcs (regulator of capsule
synthesis) two-component system is the master regulator of EPS synthesis
in enterics (Leigh and Coplin, 1992). Positive regulators homologous to
E. coli rcsA and the two-component rcsB/C were cloned by
complementation of the corresponding E. coli mutants with P. stewartii
cosmids (Poetter and Coplin, 1991). An RcsA/RcsB heterodimer was later
shown to specifically bind a 23-bp fragment upstream of the cpsA gene of
P. stewartii (Wehland et al., 1999).
EPS production and virulence is also controlled by an acyl-homoserine-
lactone-based quorum sensing system (von Bodman and Farrand, 1995);
reviewed by (Von Bodman et al., 2003) comprising the EsaR/EsaI proteins
(Vibrio fisherii LuxR and LuxI homologs, respectively) (Fig. 1.3). These
29 proteins are encoded by two convergently transcribed genes and
expression of esaR is autoinducible. A putative lux box-like promoter
element is present in the –10 region of the esaR gene (von Bodman and
Farrand, 1995). The major AHL was detected in culture supernatants using
an Agrobacterium tumefaciens NT1(pJM749, pSVB33) reporter strain.
This was purified and identified as N-(-3-oxohexanoyl)-L-homoserine
lactone (OHHL) by electrospray fast atom bombardment mass
spectrometry (von Bodman and Farrand, 1995). In addition, thin layer
chromatography later revealed the production of subnanomolar amounts of
N-(-3-oxooctanoyl)-L-homoserine lactone (OOHL). Expression of esaI is
constitutive (von Bodman and Farrand, 1995) and production of OHHL is
linear with growth, whereas OOHL was detected only at high cell
densities (Von Bodman et al., 1998). The crystal structure of EsaI was
recently solved at a resolution of 1.8 Å (Watson et al., 2002). Catalytic site modeling and site-directed mutagenesis allowed the authors to define critical residues for catalytic activity and acyl-chain specificity involved in the acylation reaction with S-adenosyl-L-methionine (Watson et al.,
2002). EPS production and virulence (using a pin-prick leaf inoculation assay) was abolished in an OHHL synthase (esaI) mutant (von Bodman and Farrand, 1995). This effect was suppressed either by a mutation in the esaR locus or addition of exogenous OHHL. Production of EPS by the esaR and esaRI null mutants was constitutive. Together these results suggested a novel negative regulatory role for EsaR at low concentrations
30 of OHHL (i.e. low cell densities) (Von Bodman et al., 1998). EsaR was later shown to bind to the esaR promoter in the absence of OHHL and
increasingly higher concentrations of OHHL titrated out EsaR from the
protein-target DNA complex (Minogue et al., 2002). No direct binding of
EsaR to any cps promoter or other target genes has yet been reported, so it
is therefore conceivable that esaR may act through an intermediate
regulator(s).
Another global regulatory system controlling virulence traits is the
GacA/GacS two-component system (Heeb and Haas, 2001). GacA belongs to the FixJ class of response regulators and GacS has a domain organization typical of unorthodox sensor kinases (Tsuzuki et al., 1995;
Stock et al., 2000) (See section 1.3.2). First identified in Pseudomonas syringae pv. syringae as a gene required for swarming, lesion formation, and protease and toxin production (LemA=GacA) (Barta et al., 1992;
Hrabak and Willis, 1992; Rich et al., 1992; Rich et al., 1994; Rich and
Willis, 1997; Kinscherf and Willis, 1999), this two-component regulatory system was found to be conserved in many gamma proteobacteria.
GacA/GacS is involved in soft rot development in P. marginalis (Liao et al., 1997), antibiotic and exoenzyme synthesis in Pseudomonas fluorescens (Corbell and Loper, 1995; Whistler et al., 1998; Bull et al.,
2001; Mascher et al., 2002; Zuber et al., 2003), phenazine production in
Pseudomonas aureofaciens (Chancey et al., 1999; Whistler and Pierson,
2003), exoenzyme and siderophore production in Pseudomonas viridiflava
31 (GacA=RepB) and P. marginalis (Liao et al., 1997), biofilm formation
and rpoS expression in P. aeruginosa (Reimmann et al., 1997; Tan et al.,
1999; Parkins et al., 2001), modulation of the ToxR regulon in Vibrio
cholerae (Wong et al., 1998), virulence, exoenzyme and harpin production
in P. carotovorum (Frederick et al., 1997; Eriksson et al., 1998; Cui et
al., 2001; Hyytiainen et al., 2001), alginate and polyhydroxybutyrate
production in Azotobacter vinelandii (Castaneda et al., 2000), perception
of intestinal signals and virulence gene activation in Salmonella enterica
serovar Typhimurium (BarA/SirA) (Johnston et al., 1996; Ahmer et al.,
1999; Prouty and Gunn, 2000; Goodier and Ahmer, 2001; Lawhon et al.,
2002), and switching between glycolytic and gluconeogenic carbon
sources, rpoS expression, and biofilm formation in E. coli (BarA/UvrY)
(Mukhopadhyay et al., 2000; Pernestig et al., 2001; Pernestig et al.,
2003). In P. carotovorum, GacA/GacS(=RpfA) controls transcription of
rsmB (homologous to E. coli csrB), a small regulatory RNA that post-
transcriptionally controls RsmA (homologous to E. coli CsrA) expression
(Cui et al., 2001; Chatterjee et al., 2002). A similar regulatory pathway has been described in E. coli, with UvrY directly controlling csrB transcription in a csrA mutant background (Suzuki et al., 2002). Southern blot hybridizations (Cui et al., 2001) demonstrated the presence of gacA
and gacS homologs in several non-soft-rotting Erwinia/Pantoea species, such as E. amylovora, E. rhapontici, P. agglomerans, P. agglomerans pv.
gypsophilae and P. stewartii, The phenotype of gacA/gacS mutations in P.
32 stewartii is not known, but these genes are likely to play a role in regulating virulence similar to that described in P. carotovorum and P.
syringae.
The most important mechanism of virulence of P. stewartii identified so
far is represented by the expression of a type III secretion (TTS) system
that secretes anti-host effector proteins into the plant apoplast (i.e the
intercellular spaces and the cell walls) or translocates them into the
symplast (i.e. the cytoplasm) (Discussed in section 1.2). Phytobacterial
type III secretion systems are called Hrp (for Hypersensitive response and pathogenicity and pronounced “harp”). These are conserved in many
gram-negative plant pathogens, including P. stewartii, which has a typical hrp cluster that spans over 40-kb. It is required for HR elicitation in tobacco and disease in corn (Coplin et al., 1986; Coplin et al., 1992;
Frederick et al., 1993; Ahmad et al., 2001; Frederick et al., 2001).
Expression of this system is essential for active colonization of the
apoplast and growth to high population densities (>106 CFU/g of wet
tissue). The other major virulence factor, EPS, is only required for
systemic spread, wilting and maintenance of water-soaking, since EPS
mutants with a functional Hrp TTS system can still colonize the
intercellular spaces of leaves, the xylem and cause initial water-soaking.
33
1.2 TYPE III SECRETION SYSTEMS, HRP GENES AND THEIR
REGULATION
1.2.1 Bacterial secretion systems and the type III secretion apparatus
An impressive number of reviews and book chapters are available on the
topic of bacterial protein secretion in general and type III secretion in particular (Cornelis and Wolf-Watz, 1997; He, 1997; Cornelis, 1998;
Hueck, 1998; Galan and Collmer, 1999; Kaper and Hacker, 1999; Cossart
et al., 2000; Koster et al., 2000; Christie, 2001; Sandkvist, 2001;
Cornelis, 2002; Fischer et al., 2002; Wilson et al., 2002). In this section I
will briefly describe both topics. My objective is to provide a background
for later discussion of phytobacterial Hrp/type III secretion systems and
their regulation, while remanding the reader to the literature for more
details on specific systems (Beer et al., 1991; Hutcheson, 1999; Collmer
et al., 2000; Cornelis and Van Gijsegem, 2000).
Secretion of proteins through membranes is one of the most important
activities of bacterial and eukaryotic cells (Cossart et al., 2000). In
bacteria, it is estimated that about 20% of the proteins synthesized are
targeted outside the cytoplasm (Wilson et al., 2002). Bacteria need
protein secretion to synthesize extracellular structures -such as flagella,
pili and fimbriae-, to degrade and transport into the cell various classes of
macromolecules, to build functional membranes, and to target toxins to
34 the extracellular or intracellular compartments of eukaryotic cells, while they are acting as partners in symbiotic or parasitic interactions.
Gram-negative bacteria have several compartments that include the cytosol, the cytoplasmic membrane, the periplasm and the outer membrane. Secreted proteins synthesized in the cytoplasm may be targeted to another compartment or to the extracellular environment via various pathways. Known secretory pathways include: i) the type I secretion systems, which are part of the complex family of ABC transporters, ii) the twin-arginine translocation (Tat) pathway, iii) the type III secretion systems, and iv) the general secretory pathway (GSP), with its four terminal branches, namely the chaperone-usher system, type
II secretion pathway, type IV secretion and type V autotransporters (Fig.
1.4) (Wilson et al., 2002).
Type I secretion system substrates do not have N-terminal cleavable signal sequences, but instead possess targeting sequences in their C- terminal domains. Type I apparatuses are composed of three proteins and are energized by ATP hydrolysis, as are all the other secretion systems.
Substrates of such widespread systems include the RTX haemolysin toxin of E. coli, antibacterial peptides (such as bacteriocins and antibiotics), and the Bordetella pertussis cyclolysin toxin. Proteins traveling the general secretory pathway in gram-negative bacteria have an 18- to 26- amino acid N-terminal signal sequence that is cleaved by a signal peptidase during secretion across the inner membrane. This pathway is
35 exemplified by the Sec system of E. coli, which is composed of a Sec
translocase complex located on the cytoplasmic membrane and one or
more aspecific cytoplasmic chaperones (such as SecB, DnaK, DnaJ or
GrpE), which keep the substrate unfolded. The GSP only translocates its
substrates to the periplasmic compartment. In many cases this is not the
final destination, so that other secretion pathways, collectively called
“terminal branches” of the GSP, are involved in translocation across the outer membrane. The main terminal branch (MTB) or type II secretion is the most commonly used pathway in gram-negative bacteria. This system is composed of 12 to 14 proteins, including an outer membrane multimeric
(10-12 subunits) secretin (e.g. Klebsiella oxytoca PulD), which has a
central pore large enough (90 Å in diameter) to accomodate folded
proteins (Wilson et al., 2002). Type II secretion is involved in the export
of many enzymes involved in degradation of macromolecules and/or in
virulence. Examples include the P. aeruginosa phospholipase C, alkaline
phosphatase, and endotoxin A (via the xcp-endoded system), Vibrio
cholerae Cholera Toxin, Type IV pili involved in twitching motility and adhesion of many gram-negative pathogens, pectic enzymes and cellulases though the P. carotovorum out system, and polygalactorunases via the
Xanthomonas campestris xps-encoded system (reviewed in (Hobbs and
Mattick, 1993). The chaperone-usher pathway is involved in the secretion of specific external appendices, such as pili and fimbriae. The type V systems involve protein substrates, whose C-terminal portions include a
36 pore-forming domain, which is able to fold into a 14-strand beta-barrel structure in the outer membrane that directs the translocation of the N- terminal “passenger” domain (Wilson et al., 2002). Type IV pathways secrete either proteins or nucleoprotein complexes. In at least two cases, type IV systems vectorially translocate DNA and/or proteins into eukaryotic cells. The first case is exemplified by the vir system of
Agrobacterium tumefaciens, which uses a dedicated inner membrane
secretion machine as an alternative to the GSP. The second example is the
cag system of Helicobacter pilori (Backert et al., 2002). Other examples
of substrates traveling this pathway are Bordetella pertussis Pertussis
Toxin and the Legionella pneumophila LcmX toxin (Fischer et al., 2002).
Type IV systems are thought to have evolved from plasmid conjugation systems (Christie, 2001; Fischer et al., 2002). The twin-arginine secretion system has been recently described as a distinct general secretory pathway that recognizes specific, cleaved targeting signals and is able to secrete fully folded substrates (Wilson et al., 2002).
TTS systems are unique in that they always mediate secretion/translocation of proteins not just across bacterial membranes, but also across eukaryotic membranes (cytoplasmic and/or phagosomal) in a vectorial fashion. In this respect, they truly function as molecular syringes injecting proteins into a eukaryotic host cell’s cytosol. These systems are always found in bacteria that spend some part of their life cycle in close interaction with either plant or animal cells. The bacterial
37 proteins injected into the host cells act as “powerful effectors” able to
subvert the host’s physiology to the bacterium’s advantage (Cornelis,
2002), thereby initiating “biochemical cross-talk” between the two bionts
(as defined by J. E. Galan; Galan and Bliska, 1996). Substrates of TTS systems do not have cleavable N-terminal sequences and the secretion step is believed to occur without periplasmic intermediates, with the exception of P. syringae (Charkowski et al., 1997). The nature of the secretion signal is still controversial and N-terminal polypeptide amphipathicity/primary sequence composition (Lloyd et al., 2001; Lloyd et al., 2002) a 5’-mRNA signal (Anderson and Schneewind, 1997;
Anderson et al., 1999; Anderson and Schneewind, 1999; Hienonen et al.,
2002) and/or chaperone-mediated targeting (Cheng and Schneewind,
1999), have been shown in various systems.
TTS systems were described for the first time in Yersinia enterocolitica,
Yersinia pseudotubercolosis and Yersinia pestis between 1991 and 1994
(Michiels and Cornelis, 1991; Michiels et al., 1991; Allaoui et al., 1994;
Sory and Cornelis, 1994; Woestyn et al., 1994). Since then, more than 20
TTS systems have been reported in animal, plant and insect pathogens and symbionts (Table 1.1). Besides Yersinia spp. (reviewed by (Cornelis,
2002), TTS apparatuses have been described in many other gram-negative animal pathogens, such as Salmonella spp. (reviewed in (Galan and
Collmer, 1999), Shigella spp. (Van Nhieu, 1997), enteropathogenic E.coli
(EPEC) and enterohemorragic E. coli (Frankel et al., 1998; Torres and
38 Kaper, 2002), P. aeruginosa, Chlamydia trachomatis, Chlamydia psittaci,
Chlamydia pneumoniae, Bordetella bronchiseptica, Bordetella pertussis,
and Burkholderia pseudomallei. The contribution of the TTS system to the
disease cycles of these pathogens at the cellular level varies and often
spans opposite behaviors in different species, ranging from inhibiting
phagocytic cells (e.g. in Yersinia spp.) to stimulating internalization by
nonphagocytic cells (in Salmonella enterica and Shigella spp.). In general, the net effect is to favor entry or survival of the invading pathogen by disrupting the host metabolism or disrupting intracellular signal transduction pathways, thereby causing a variety of symptoms, such as mild/severe diarrhea, septicemia, septic shock, and chronic infections
of the lung (Muller et al., 2001). TTS systems have also been described in
all gram-negative, non-tumorigenic phytobacteria surveyed so far, namely
P. syringae pathovars (1985-86) (Lindgren et al., 1986), P. stewartii
(Coplin et al., 1986), P. agglomerans pv. gypsophilae (Nizan et al.,
1997), E. amylovora (Steinberger and Beer, 1988; Barny et al., 1990; Beer
et al., 1991), P. carotovorum (Bell et al., 2002), P. chrysanthemi (Bauer
et al., 1994), X. campestris pv. campestris (Arlat et al., 1991) and pv.
vesicatoria (Arlat et al., 1991) and Ralstonia solanacearum (Boucher et
al., 1987). TTS apparatuses in phytobacteria are encoded by a subset of
the so-called hrp genes, which are discussed in greater detail in the next
section. TTS systems have also been described in the plant symbiont
Rhizobium leguminosarum (Viprey et al., 1998; Perret et al., 1999; Krause
39 et al., 2002) the plant growth promoting P. fluorescens strain SBW25
(Preston et al., 2001) and in the insect symbiont Sodalis glossinidius and
the entomopathogenic bacterium Photorhabdus luminescens (Ffrench-
Constant et al., 2000; Dale et al., 2001). In phytobacterial pathogens, TTS systems promote bacterial growth in plant tissues and subsequent disease symptom development, especially water-soaking or edema-like lesions and necrosis. They are also required for elicitation of rapid programmed defense responses that are induced in resistant plants in response to the specific elicitor proteins injected by the invading bacteria. These then trigger plant cell death, thereby restricting the infection (Staskawicz et al., 2001).
By far, the best-characterized TTS system is the Ysc (Yop secretion) apparatus of Yersinia spp. (Cornelis, 2002), also called the Ysc injectisome. The proteins traveling this system are called Yops (for
Yersinia outer proteins). The Ysc apparatus is an organelle made of a basal body spanning the inner membrane, the peptidoglycan layer and the outer membrane. The basal body is extended toward the external environment with a rigid needle-like structure. The basal body architecture resembles the flagellum basal body (Fig. 1.5), with the proximal part of the structure being formed by a large cylinder (YscQ) and an inner membrane-spanning ring (YscJ), corresponding to the flagellar C and MS rings, respectively. This portion also contains the protein-pumping apparatus (analogous to the flagellum-specific secretion
40 apparatus) comprised of YscV, YscU, YscR, YscT, YscS and the ATPase motor YscN. The distal part of the body, which does not have homologous proteins in the remaining structure of the flagellar apparatus, forms a 200-
Å ring-shaped structure with a hollow center of about 50 Å in diameter that is made of YscC monomers. YscC belongs to the PulD/pIV superfamily of outer membrane secretins. This secretin-like multimer is probably stabilized by the lipoprotein YscW. The outer ring is most likely connected to the basal body proximal cylinder by the lipoprotein YscJ.
However, YscJ has not yet been localized in Yersinia spp. (Cornelis,
2002). The Ysc needle is a rigid polymeric rod about 1200 Å long and 70
Å wide, mainly formed by the 6-kDa YscF monomer and three other proteins and containing a hollow conduit 20 Å in diameter. This needle- like structure can be isolated by physical methods and is usually associated with the basal body secretin-like protein and YscJ. The injectisome can release Yops into the external environment, but that alone is not sufficient for the translocation of proteins into eukaryotic cells.
Three additional Yop translocators (YopB, YopD and LcrV) are required for this step. Therefore, the description of the injectosome as a syringe directly poking holes into eukaryotic membranes is somewhat misleading
(Cornelis, 2002). An important property of TTS machines is that, at least in some animal pathogens (Yersinia spp., P. aeruginosa and Shigella spp.), release of TTS substrates is dependent on contact with host cell surfaces, which in some systems may be mimicked in vitro by modulating
41 divalent cation concentration and temperature (e.g. Yersinia spp.) or by
adding amphipathic dyes, such as Congo Red, or serum proteins (e.g.
Shigella spp.) (Francis et al., 2002). In addition, cell contact also
increases TTS system gene transcription upon secretion of negative
regulators, such as LcrQ (Petterson et al., 1996). For one plant pathogen, cell-contact regulation of the hrp-TTS genes has been suggested (Aldon et
al., 2000).
1.2.2. Phytobacterial Hrp type III secretion systems
Many reviews on the discovery of hrp genes and their structural and
functional characterization have been published in the last decade (Willis
et al., 1991; Bonas, 1994; van Gijsegem et al., 1995; Alfano and Collmer,
1997; Lindgren, 1997; He, 1998; Hutcheson, 1999; Cornelis and Van
Gijsegem, 2000). More recently the emphasis in this area has shifted to the functional analysis of substrates of the Hrp Type III secretion system
(such as Avr proteins and other effectors) in various host-pathogen model systems (Bonas and Van den Ackerveken, 1999; Buttner and Bonas,
2002). In this section, I will present a brief overview of the properties of this class of secretion machines and their substrates.
hrp genes were first identified in P. syringae pv. syringae in 1985-1986
(Lindgren et al., 1986), well before the identification of mammalian pathogen TTS systems. Only in 1992, did studies on the DNA sequences of hrp genes from X. campestris pv. vesicatoria (Fenselau et al., 1992)
42 and R. solanacearum (Gough et al., 1992) first reveal homologies to some of the proteins in TTS systems of Yersinia spp. and Shigella flexneri.
Initially, hrp genes were functionally defined based on their mutant phenotype; hrp genes had to be dispensable for growth in minimal media, but essential for pathogenicity in susceptible hosts and for eliciting the hypersensitive response (HR) in incompatible hosts and non-hosts
(Lindgren et al., 1986). The HR is a rapid and localized defense response of the plant that involves localized programmed cell death, active oxygen species production, and synthesis of phenolics and antimicrobial compounds at and around the site of infection (Klement, 1982; Klement et al., 2003). During natural infections, the reaction is usually not visible because too few host and bacterial cells are involved, but, under experimental conditions, high doses of inoculum (>107 /ml) will cause
massive cell collapse and confluent necrosis that is visible to the naked
eye (Klement, 1982). This reaction has the net effect of restricting
pathogen multiplication and further progress of the disease. HR is not
elicited by true non-pathogens, including many saprophytic bacteria and
the molecular cloning host E. coli, even at high inoculum doses.
hrp genes are clustered, spanning up to 41 kb of DNA (van Gijsegem et
al., 1995) and are located either on the chromosome or large plasmids
(Boucher et al., 1987). In some cases, cosmid clones containing nearly
complete hrp clusters enable non-pathogenic bacteria, such as E. coli K12
and P. fluorescens, to cause an HR, but not disease (Lindgren, 1997).
43 Subsequent DNA sequencing of overlapping cosmid clones spanning hrp
clusters and mutational analysis showed that hrp clusters encoded not
only the proteins involved in secretion, but also proteins involved in the
specific regulation of hrp and effector genes, elicitors of the HR in non-
host plants, and disease-specific (Dsp) proteins involved in pathogenicity
but not HR elicitation in Erwinia/Pantoea spp. (Bogdanove et al., 1998;
Bogdanove et al., 1998; Mor et al., 2001). Sequencing of the flanking
regions of hrp clusters (especially Pseudomonas syringae; (Alfano et al.,
2000; Collmer et al., 2000; Noel et al., 2002; Deng et al., 2003) and the completion of several genome projects (Bell et al., 2002; da Silva et al.,
2002; Salanoubat et al., 2002; Buell et al., 2003) have revealed that sets of putative substrates of the Hrp secretion apparatus are commonly located nearby. Another realization from these studies is the concept of hrp clusters as horizontally mobile pathogenicity islands (PAIs)
(Hutcheson, 1999). The presence of remnants of transposons and insertion sequences and, in some cases, tRNA genes (Alfano et al., 2000) at the extreme ends of the hrp/effector cluster is indicative of previous mobilization via plasmids, conjugative transposons or phages followed by integration into the chromosome or plasmids of the recipient strain (Kaper and Hacker, 1999). Homing of the PAI during the evolution of the species supposedly degraded its ability to mobilize further (Kaper and Hacker,
44 1999). In some species, a few loci showing Hrp phenotypes were also
found to be unlinked to the major hrp cluster, e.g. Xanthomonas hrpX and
P. syringae hrpM (Reviewed in van Gijsegem et al., 1995).
Phytobacterial TTS systems are usually encoded by three to four operons for a total of 19 to 21 genes (Hutcheson, 1999). Up to 12 of the encoded proteins show similarities with Ysc proteins that represent the core components of the secretion apparatus. Of these, eight share similarities with flagellum apparatus proteins and nine are conserved among all phytobacterial TTS systems (He, 1997; He, 1998). The nine conserved genes have been named hrc, which stands for hrp conserved
(Bogdanove et al., 1996), and each gene uses the same fourth letter as its
corresponding ysc homolog.
hrp gene clusters have been classified into two groups based on their genetic organization and manner of regulation (Alfano and Collmer, 1997)
(Hutcheson, 1999). Group I hrp clusters include those initially described in P. syringae pathovars, reviewed in (Hutcheson, 1999) and (Collmer et
al., 2000), and Erwinia amylovora (Steinberger and Beer, 1988; Beer et
al., 1991; Laby and Beer, 1992; Wei et al., 1992; Wei and Beer, 1993;
Wei and Beer, 1995; Kim et al., 1997; Kim and Beer, 1998; Wei et al.,
2000b). Later studies found nearly identical clusters throughout the
erwinias (Bauer et al., 1994; Frederick et al., 2001; Mor et al., 2001) and
NCBI unfinished microbial genomes). In erwinias, the hrpJ (containing
hrcV-hrcN), hrpU (containing hrcQRSTU), hrpA (containing hrcJ) and
45 hrpC (containing hrcC) operons are collinear with the corresponding transcriptional units in P. syringae, with the minor difference that the hrpU-J operon is split in two different transcriptional units (Fig. 1.6)
(Huang et al., 1993; Lidell and Hutcheson, 1994; Preston et al., 1995;
Kim et al., 1997; Deng et al., 1998; Kim et al., 1998). In erwinia hrp clusters, the hrpA-hrpC operons and hrpJ-hrpU operons are divergently transcribed with the master regulatory genes (hrpL, hrpXY and hrpS) located in between (Bauer et al., 1994; Wei et al., 2000b; Frederick et al.,
2001; Nizan-Koren et al., 2003). In P. syringae, the above operons are convergently transcribed and the regulatory genes are located on each end
(hrpL is on one side and hrpRS on the other). Unlinked candidate orthologs for hrpX and hrpY have been found in P. syringae and in P. aeruginosa, but their role is not known. Many TTS system effector genes are located in the regions flanking the core cluster (Fig. 1.6). These strong similarities suggest horizontal transfer and subsequent homing- related recombination/transposition events (Hutcheson, 1999).
The second group of hrp clusters is found in Xanthomonas spp. and R. solanacearum. These group II clusters are basically collinear to each other, with the only difference being the position of the regulatory genes hrpXXc vs. hrpBRsol (Arlat et al., 1991; Van Gijsegem et al., 1995). The
secretion genes are organized into four operons; hrcC, hrcJNT, hrcUV and hrcQRS, the last two being transcribed in the opposite direction. The sequence similarity of group II hrc genes to their mammalian TTS system
46 homologs is less than that found for group I hrc genes (Hutcheson, 1999).
The regulatory systems for group I and II hrp gene are completely different; group I hrp cluster are regulated by the alternative sigma factor
HrpL, whereas group II are regulated by AraC-like transcriptional activators (HrpB for R. solanacearum and HrpX for Xanthomonas spp.).
Further details are discussed below.
The proteins secreted by the hrp/hrc TTS systems have been named
Hops (for Hrp outer proteins), to parallel the designation of Yersinia outer proteins as Yops (Alfano and Collmer, 1997). At least four classes of
Hops travel the secretion apparatus: Harpins, Hrp pilins, translocators and effector proteins. Harpins, Hrp pilins and translocators are collectively defined as helper proteins for their presumed or demonstrated roles in aiding the secretion/translocation of other TTS system substrates (Collmer
et al., 2002).
Harpins are hydrophilic, glycine-rich, heat-stable, cysteine-free proteins
that are able to elicit an HR when infiltrated at high concentration into the
plant apoplast (Alfano and Collmer, 1997). Harpins travel through the
TTS apparatus but are released into the apoplastic spaces, rather than
being injected into host cells (Wei et al., 1992; Bogdanove et al., 1996;
Pike et al., 1998; Perino et al., 1999). Harpins have been described in E.
amylovora (harpinEa or HrpN and HrpW; Wei et al., 1992; Gaudriault et
al., 1998; Kim and Beer, 1998) in other erwinias (Bauer et al., 1995;
Mukherjee et al., 1997; Ahmad et al., 2001), in all P. syringae pathovars
47 surveyed (harpinPss or HrpZ; He et al., 1993; Charkowski et al., 1998), and in R. solanacearum (PopA1; Arlat et al., 1994), but no sequence similarity exists among the HrpN, HrpW, HrpZ and PopA1 proteins. The precise role of harpins in pathogenesis is currently unknown. In general, they are abundantly secreted into the growth medium and therefore were the first Hops to be characterized. Initially harpins were thought to be the universal HR elicitor molecule (Wei et al., 1992; Beer et al., 1993; He et al., 1993; Bauer et al., 1995), but subsequent studies in Pseudomonas syringae questioned this conclusion (Alfano et al., 1996). In P. stewartii and E. amylovora, hrpN mutations greatly affect HR elicitation in tobacco but not virulence (Barny, 1995; Ahmad et al., 2001). The reason for these differences in phenotypes could be due to functional redundancy in P. syringae or to differences in the secretion mechanism. HrpZ is able to bind plant cell walls (Hoyos and al., 1996) and create transient pores in lipid bilayers (Lee et al., 2001). P. syringae and Erwinia amylovora HrpW proteins have a C-terminal pectate lyase domain that binds pectin, a major component of cell walls (Charkowski et al., 1998; Kim and Beer, 1998).
These properties may play a role in facilitating translocation of effectors
(Collmer et al., 2002) or in the release of nutrients from host cells (Lee et al., 2001). Harpins are also of commercial interest because they induce systemic resistance responses when sprayed on plants (Dong et al., 1999;
Kariola et al., 2003).
48 Hrp pilins, such as P. syringae HrpA (Roine et al., 1997), E. amylovora
HrpA (Wei et al., 2000a), X. campestris HrpE1 (He and Jin, 2003), R.
solanacearum HrpY, (Van Gijsegem et al., 2000) are small proteins abundantly secreted into the growth medium (for a recent review see He and Jin, 2003). In contrast to animal pathogens (Knutton, 1998), Hrp pili are essential for secretion of other Hops (Van Gijsegem et al., 2000; Wei et al., 2000a). P. syringae HrpA has been shown to spontaneously assemble into a pilus-like structure in vitro (Roine et al., 1997). Unlike the short rigid needles of mammalian pathogens, Hrp pili have been observed in vivo as flexuous filaments of ca. 80 Å x 2 µm. They appear to be extended by condensation of monomers at the distal end (Jin and He,
2001). Hrp pilus assembly in vivo is hrp/hrc dependent. Hrp pili have been shown to cross the thick (100-200 nm) plant cell wall (Hu et al.,
2001).
Xanthomonas HrpF is the only Hop shown by genetic and biophysical approaches to behave as a putative translocator (Buttner et al., 2002).
HrpF is a predominantly hydrophilic protein with two hydrophobic domains at the C-terminus. It is highly similar to Rhizobium fredii NolX, which is required for legume host specificity (Huguet and Bonas, 1997).
Non-polar mutations in hrpF produce a Hrp phenotype but do not alter secretion of other Hops in vitro (Rossier et al., 2000). Experiments with lipid bilayers showed binding of HrpF and HrpF-dependent pore formation
(Buttner et al., 2002).
49 The fourth group of proteins traveling the Hrp secretion apparatus
encompasses the Hop-effectors. This is by far the largest class of Hops,
comprising more than 30 proteins per species in some cases (Collmer et
al., 2002). Secretion levels of Hops in culture and in planta are much
lower than for harpins or Hrp pilins. This delayed their discovery as TTS
system substrates. All Hop effectors share a few common properties. They
have no effect on the plant cell when applied to the apoplast and they are
vectorially delivered into the host cell. This has been shown by direct
approaches using immunocytochemical and biochemical assays, e.g.
AvrBs2 (Casper-Lindley et al., 2002) and AvrBs3 (Szurek et al., 2002), or by indirect approaches using transgenic expression and/or two-hybrid analysis (Gopalan et al., 1996; Van den Ackerveken et al., 1996; Mudgett
and Staskawicz, 1999; Kim et al., 2002). In most cases, mutagenesis of
hop genes has either no effect or only quantitatively affects virulence
(Ritter and Dangl, 1995; Alfano et al., 2000; Lavie et al., 2002; Deng et
al., 2003), suggesting functional redundancy or additivity. Some effectors
have functional nuclear localization signals and/or features of eukaryotic
transcription factors (Gueneron et al., 2000; Szurek et al., 2001; Marois
et al., 2002; Deslandes et al., 2003). Therefore, they are presumed to act
by directly altering gene expression patterns of the host. Enzymatic
activities have been shown or proposed for a few of them, as discussed
below.
50 Three subclasses of Hop effectors have been recognized from an
operational viewpoint: Avr (avirulence) proteins, Dsp/Pth (disease
specificity/pathogenesis) proteins and candidate Hop effectors. This
division is somewhat artificial, because at a molecular level their
mechanisms of action probably overlap in many cases, but it reflects the
historical and practical approach still found in the literature.
Avr (avirulence) proteins are effectors that are recognized by a resistant
plant’s surveillance system, which is represented by R gene products and
other accessory proteins (reviewed by McDowell and Dangl, 2000; Dangl
and Jones, 2001; Staskawicz et al., 2001; Holt et al., 2003). Their existence was predicted by the gene-for-gene hypothesis elaborated during the late 1930s-early 1940’s (reviewed by Flor, 1971). avr genes in bacteria were initially cloned by screening collections of virulent strains
(able to cause disease on compatible hosts) carrying cosmid libraries established from avirulent strains (able to cause HR on incompatible hosts). These transcojugants were assayed for gain of incompatibility phenotype on host cultivars carrying specific R genes (Staskawicz et al.,
1984; Gabriel et al., 1986; Staskawicz et al., 1987). Mutations in this subclass of effectors result in loss of HR elicitation in resistant/incompatible cultivars (i.e. expressing cognate R proteins) of a given host species, but usually they have no major effects on disease development in susceptible/compatible cultivars (i.e. lacking the cognate
R gene). Avr proteins are a result of the coevolution of pathogens with
51 their host resistance gene system. Genes encoding this class of proteins are frequently found in Xanthomonas and Pseudomonas, two genera of phytobacteria able to infect a great variety of hosts. Typically these species have several dozens of hop genes per genome. For some Avr proteins a great deal of information on their interaction with the R-protein system is available, but their primary function in virulence is often unknown. In a few cases, clear effects on virulence are observed.
Transposon mutations in P. syringae avrPto, avrE and avrRpm1 and
Xanthomonas avrBs2 caused a dramatic reduction of growth in planta and symptom development in specific hosts (Ritter and Dangl, 1995; Swords et al., 1996; Chang et al., 2000). Suppression of programmed cell death is another function that has been attributed to Avr/Hop effectors. Evidence for this often comes from studies where transgenic expression of Avr proteins is susceptible genetic backgrounds. For instance, AvrPtoB suppresses Pto-AvrPto-based HR defense responses leading to increased susceptibility (Abramovitch et al., 2003); similarly, cell wall based defenses (e.g. callose depositions) were suppressed in a salicylic acid- dependent manner by ectopic expression of AvrPto in Arabidopsis thaliana (Hauck et al., 2003).
Dsp/Pth proteins are operationally defined as effector proteins that contribute to virulence and symptom development for which no cognate
R-gene has been identified. One subgroup of Dsp proteins, which have high sequence similarity, has been described in erwinias. It includes E.
52 amylovora DspA/E, P. stewartii WtsE, P. agglomerans pv. gypsophilae
DspE (Gaudriault et al., 1997; Frederick et al., 2001; Mor et al., 2001) and P. carotovorum DspE (Yang et al., 2002). These all share amino acid similarity to P. syringae AvrE (Lorang and Keen, 1995). A second example of avr-like genes that are essential for virulence is X. citri PthA, a member of the large avrBs3 family (Dangl, 1994). pthA expression in X. campestris changes the pathogen’s host-specificity (Swarup et al., 1991).
Mutations in pthA eliminate virulence on its normal host citrus (Swarup et al., 1992), and transient expression of pthA in susceptible host cells led to elicitation of cell death and hypertrophic symptoms (Duan et al., 1999).
Candidate effectors are putative Hops for which a virulence or avirulence function has not been demonstrated. They have been identified by sequence homology to known Hops or by various screening procedures.
These include in silico screens of genomic sequences for Hrp promoters and secretion motifs or functional screens for Hrp regulated promoters,
Hrp-dependent secretion or in planta expression (Boch et al., 2002; Fouts et al., 2002; Petnicki-Ocwieja et al., 2002), reviewed in (Collmer et al.,
2002). They are the fastest growing class of effectors. Some Hop effectors are homologous to known avirulence proteins. For example, HopPtoJ is the P. syringae homolog of X. vesicatoria AvrXv3 and HopPtoC is the homolog of AvrPpiC2 (Petnicki-Ocwieja et al., 2002). For some, Hrp- dependent secretion and or translocation has been demonstrated (e.g.
HopPtoD1, HopPtoE, HopPtoG, HopPtoH, HopPtoJ, HopPtoK and many
53 others, Petnicki-Ocwieja et al., 2002; HopPtoB1, Alfano et al., 2000;
Deng et al., 2003; HopPtoA1 and HopPtoA2, Badel et al., 2002). Others have enzymatic properties that enable them to modify plant defense responses. For example, HopPtoD2 is a tyrosine phosphatase (Espinosa et al., 2003) and AvrPphB is a Cys-protease (Shao et al., 2002). Sequence homology suggests that enzymatic activities may be associated with other
Hops. For example, HopPmaG may be a transglycosylase; HopPmaH may be a pectin lyase; HopPtoC and HopPtoN may be cysteine-proteases; and
HopPtoS3 may be an ADP-ribosyltransferase (reviewed in Collmer et al.,
2002). Recently, another candidate hop gene, HopPtoA1, has been shown to contribute to P. syringae colony formation in the apoplast of a compatible host (Badel et al., 2002).
1.2.3 Regulation of type III secretion systems
During the infection process, TTS systems are transcriptionally and post-translationally activated in a highly regulated fashion. The pathogen responds to initial cues provided by the host, adapts to subsequent host responses, and thereby creates a new niche while modulating its “invasion program” over time and space. In mammalian pathogens, two distinct steps can be recognized (Francis et al., 2002): an initial phase where components of the TTS system are made and assembled and then a second phase where the effector proteins are produced and targeted for secretion.
In reality, some temporal overlapping occurs, which may contribute to
54 differences in the timing and ordered secretion of various classes of effectors. In plant pathogens, the secretion apparatus and the effector genes always appear to be co-regulated at the transcriptional level. In general, one important difference between animal and plant pathogens is that gram-negative plant pathogens always remain extracellular. By adhering to the cell wall, some 100-200 Å away from the host plasma membrane, they experience a relatively homogeneous environment inside the host tissues. On the other hand, a mammalian pathogen must be able to deal with a diverse range of host compartments until it reaches an appropriate site of infection. Moreover, mammalian pathogens have direct access to the host membranes and some are even able to internalize and multiply in phagosomal vescicles (e.g. Salmonella and Shigella spp.)
(Cossart et al., 2000).
All TTS systems are controlled by complex regulatory networks that are able to integrate various environmental cues, including chemical, nutritional and host signals, i.e. pH; oxygen tension; redox potential;
3- 2+ 2+ ammonium, PO4 , Mg , Ca and nucleotide concentrations; growth phase; and availability of specific nutrients (Hueck, 1998), as well as microbial signals, such as quorum sensing molecules (Sperandio et al.,
1999). The necessity to minimize energy consumption or to express the secretion system only at the right time and place are often cited as the main reasons for such underlying complexity (Hueck, 1998). Among the regulators involved in controlling the transcription of TTS system
55 apparatuses and their effectors are: two-component systems (Grimm and
Panopoulos, 1989; Bernardini et al., 1990; Behlau and Miller, 1993;
Frederick et al., 1993; Xiao et al., 1994; Bajaj et al., 1995; Grimm et al.,
1995; Nakayama and Watanabe, 1995; Johnston et al., 1996; Ochman et al., 1996; Shea et al., 1996; Wengelnik et al., 1996; Merighi et al., 2003;
Nizan-Koren et al., 2003), AraC-like activators (Sakai et al., 1986;
Yother et al., 1986; Cornelis et al., 1987; Forsberg and Wolf-Watz, 1988;
Cornelis et al., 1989; Frank and Iglewski, 1991; Genin et al., 1992;
Kaniga et al., 1994; Gomez-Duarte and Kaper, 1995; Tobe et al., 1996;
Wengelnik and Bonas, 1996), alternative sigma factors of various classes
(Wei et al., 1995; Xiao et al., 1994; Cui et al., 2002), global regulators affecting DNA topology (Cornelis et al., 1991; Hromockyj et al., 1992;
Cui et al, 2002), effector protein chaperones (Darwin et al., 2001), TTS pilus components (Wei et al., 2000a), and housekeeping proteases (Bretz et al., 2002). Table 1.1 summarizes some of the known regulators of TTS systems. Examples of complex regulatory pathways described in mammalian pathogens are shown in Fig. 1.7.
1.2.3.1 Regulation of group I hrp/hrc genes
In Group I hrp clusters, the alternative sigma factor HrpL enables the recognition and transcriptional activation of promoters containing “Hrp boxes” in their –10/-35 regions (Fellay et al., 1991) (Fig. 1.8). Hrp box promoters have been found in front of all known hrp/hrc secretion
56 operons of group I Hrp clusters (Huynh et al., 1989; Innes et al., 1993;
Salmeron and Staskawicz, 1993; Shen and Keen, 1993; Xiao and
Hutcheson, 1994; Wei and Beer, 1995; Chatterjee et al., 2002; Fouts et
al., 2002) and most, but not all (Shen and Keen, 1993), avr genes. HrpL
belongs to the ECF (extracytoplasmic function) family of sigma factors
(Lonetto, 1991) but, in contrast to other members of this family (Hughes and Mathee, 1998; Raivio and Silhavy, 2001; Helmann, 2002), no anti- sigma factor has been described.
Transcription of hrpL is under the control of a σ 54 promoter based on
three observations: i) the hrpL promoter regions always contain a putative
–12/-24 σ 54 promoter consensus sequence; ii) in P. syringae, expression of hrpL is strongly reduced in an rpoN mutant (Fellay et al., 1991;
Hendrickson et al., 2000; Hendrickson et al., 2000); and iii) expression of
P. stewartii hrp genes is reduced in an E. coli rpoN strain (Frederick et
al., 1993). In P. stewartii, a region with high A+T content and an IHF-
like consensus sequence is present just upstream of the σ 54 promoter
(Merighi et al., 2001). Similarly, A. Chatterjee and coworkers (Chatterjee
et al., 2002) found that expression of Pectobacterium carotovorum hrpL
in E. coli is dependent on himA (=IHF).
Transcription initiation and elongation at σ 54 promoters typically
requires NtrC-class of enhancer-binding proteins (Hirschman et al., 1985;
Weiss et al., 1991). In this regard, all group I hrp clusters use NtrC-like
transcriptional enhancers (Grimm and Panopoulos, 1989), such as hrpR
57 and hrpS, to activate hrpL (Rahme et al., 1991; Frederick et al., 1993;
Xiao et al., 1994). This csubclass of enhancer-binding proteins have a C-
terminal DNA binding domain carrying a helix-turn-helix domain of the
Fis-like family and a σ 54-activating ATPase domain, but they lack an N- terminal phosphorylatable receiver domain (Grimm and Panopoulos,
1989). Response regulators with this structure are expected to be constitutively proficient for binding to DNA at tandem symmetrical dyad
sequences (Klose et al., 1994), even if oligomerization, which usually
follows phosphorylation of the receiver domain, may not occur. In P.
syringae, the HrpS and HrpR orthologs have been shown to interact in vitro and both proteins are required for full activation of hrpL (Hutcheson
et al., 2001). A symmetrical dyad located 90-110 bp upstream of the σ 54-
promoter is present in P. syringae (Hutcheson, 1999) and P. stewartii
(Merighi et al., 2003). In Chapter 3, we will show that this dyad is
required for activation of hrpL by HrpS in P. stewartii .
In P. syringae, expression of hrpR, hrpS and downstream genes is
environmentally regulated. Complex amino acid sources, such as peptone,
suppress expression of hrpR, hrpS and other hrp genes in P. syringae pv.
phaseolicola (Rahme et al., 1992) and other pathovars (Xiao et al., 1992;
Xiao et al., 1994). M9 minimal medium induces hrpR and hrpS to low
levels, while growth in tobacco increases their expression about 1000-fold
(Rahme et al., 1992) in as little as 1 h post-infiltration (Xiao et al., 1992).
A modified minimal medium with optimized salts, nitrogen, carbon
58 sources and pH further increases expression over M9. Interestingly, Krebs cycle intermediates, such as succinate and citrate repress hrp genes in P. syringae, probably via catabolite repression (Huynh et al., 1989). How environmental signals are coupled to hrpRS and hrpL activation is not known.
In addition to HrpS and HrpL, the erwinias also have a two-component signal transduction system encoded by the hrpXY operon, which is located downstream of hrpL. In Erwinia amylovora, Wei et al. (2001) suggested that HrpY directly controls expression of hrpL in concert with HrpS.
Growth in an acidic, low ionic strength, low phosphate, minimal medium containing mannitol and 5 mM ammonium sulfate can induce hrp genes to levels comparable to those measured in tobacco tissues and higher than those observed in pear tissues (Wei et al., 1992). Addition of nicotinic acid, certain amino acids (His, Asp, Trp and Gly), 0.1% casamino acids, or some carbon sources (glucose and maltose) repress hrp genes (Wei et al., 1992). The nature of the primary signals transduced by the two- component system and the regulatory circuits controlling hrpS are not known. In Pectobacterium spp., where hrp genes play a marginal role in virulence (Rantakari et al., 2001), hrpS is hypoyhesized to be constitutively expressed (Chatterjee et al., 2002) and the upstream pathways of hrp regulation are not characterized. P. carovovorum hrpL appears to be regulated by RpoN, HrpS and IHF at the transcriptional
59 level and by the RsmA/rsmB system at the post-transcriptional level
(Chatterjee et al., 2002). More details on this global regulatory system are
given in the Introduction to Chapter 6.
An interesting hrp-regulatory effect has been attributed to HrpA in P.
syringae. HrpA is the major component of the Hrp pilus, In addition to
being absolutely required for secretion of two effectors, the hrpA gene
appeared to be essential for full expression of genes encoding other Hrp
regulatory, secretion, and effector proteins. In particular, hrpA-mediated
gene regulation was shown to affect levels of hrpR and hrpS mRNA (Wei
et al., 2000a). The effect of HrpA appears to be unrelated to its role in
protein secretion because it is possible to isolate HrpA missense mutants
that show reduced hrpRS expression, but exhibit normal secretion of Hops
(Wei et al., 2000a).
Most of the regulatory circuits described above involve transcriptional
activation. Only one example of negative regulation of hrp genes is
known so far. A negative feed back loop triggered by HrpL activation of
the hrpC secretion operon has been described in P. syringae (Preston et
al., 1998). This may be due to over-production of HrpV, because non-
polar mutations in hrpV resulted in increased secretion of harpin and upregulation of three secretion operon reporter gene fusions. Epistasis analysis further suggested that hrpV acts as a hrp repressor upstream of hrpRS (Preston et al., 1998).
60 The Hrp regulatory cascade is also subject to post-translational regulation. Work in S. Hutcheson’s lab has recently shown that the Lon protease has a negative regulatory effect on the expression of hrpL via post-translational degradation of HrpR (Bretz et al., 2002). P. syringae lon::Tz mutants are upregulated for hrpL transcription, exhibit enhanced
AvrPto secretion, and elicit faster plant responses. Immunoblot analysis showed that the half-life of HrpR is increased in a lon mutant under hrp- inducing conditions in vitro. The authors further suggest that transcription of hrpRS is constitutive, but this contradicts other workers’ results (Wei et al., 2000a) and may be due to differences in the techniques used to measure gene expression (Northern analysis vs. RT-PCR).
1.2.3.2 Regulation of group II hrp/hrc genes
Group II hrp clusters are regulated in response to environmental stimuli similar to those affecting expression of group I hrp genes, such as growth in minimal media. For X. campestris, minimal media with glucose and/or fructose are stimulatory, but for X. vesicatoria either a tomato conditioned medium (Schulte and Bonas, 1992) or a specialized minimal medium (XVM1 and XVM2) containing methionine, sucrose and low phosphate concentrations is necessary to activate hrp genes (Wengelnik and Bonas, 1996). For R. solanacearum, pyruvate and glutamate are the most effective carbon sources for inducing hrp genes in vitro (Arlat et al.,
1992).
61 Expression of group II hrp/hrc genes is usually controlled directly by
AraC-like regulators (Fig. 1.8). In R. solanacearum the AraC-like
activator is encoded by hrpB, which was the first member of this class of hrp regulators to be discovered (Genin et al., 1992), and in X. vesicatoria, it is encoded by hrpX. Unlike hrpBRsol, hrpXXv is not linked to the hrp
cluster (Wengelnik and Bonas, 1996). These AraC-like transcription
factors directly activate expression of most of the hrp secretion operons
by recognizing specific promoter elements (Bonas, 1994; Wengelnik and
Bonas, 1996). With the exception of avrBs3 (Knoop et al., 1991), most
avr genes in Xanthomonas are also part of the Hrp regulon (Noel et al.,
2001).
Additional regulators control expression of the AraC-like genes. In R.
solanacearum, the OmpR-like response regulator HrpGRsol was shown to
control hrpBRsol expression (Brito et al., 1999). More recent results indicate that HrpGRsol may also independently control other unrelated
functions important for vascular colonization (Vasse et al., 2000).
hrpGRsol itself is controlled by a complex regulatory cascade. The TonB- like receptor PrhA (plant regulator of hrp genes) (Marenda et al., 1998) is
located in the outer membrane and perceives cell-contact signals (Aldon
et al., 2000) but not chemical/nutritional stimuli (Marenda et al., 1998).
Upon perceiving the plant signal, PrhA activates PhrI/PhrR (Brito et al.,
2002), which is probably an ECF sigma factor/antisigma factor complex.
Mutations in these regulators delay symptom development (Brito et al.,
62 2002). The PhrI sigma factor activates transcription of the FixJ-class
response regulator PrhJ (Brito et al., 1999), which in turn activates
hrpGRsol in presence of plant cells. HrpGRsol is required both for induction
in minimal medium and in planta, whereas PrhJ was only needed for
transduction of plant signals (Brito et al., 1999). This suggests that
hrpGRsol is regulated by two independent pathways, one dependent on
plant signals (perhaps cell contact) and the other on nutritional and
physiological variables.
In X. campestris, the OmpR-class, orphan response regulator HrpGXc activates transcription of hrpXXc (Schulte and Bonas, 1992). Interestingly, activation of the hrcC secretion gene is dependent on hrpGXc, but not on
hrpXXc (Wengelnik and Bonas, 1996). Certain amino acid substitutions in
HrpGXc that constitutively activate hrp gene expression also enhance the
timing and intensity of disease symptoms (Wengelnik et al., 1999). How
HrpGXc is regulated at a post-translational level and how plant and
metabolic signals are transduced is not known.
1.3 MECHANISMS AND STRATEGIES OF TWO-COMPONENT
SYSTEM REGULATION
1.3.1 Regulation of virulence factors
Bacterial pathogens activate their virulence mechanisms in response to
external cues signaling the presence of specific host compartments. This
implies tight regulation of the corresponding virulence genes. Gene
63 expression is modulated by multiple signals, including salt concentration,
pH, redox potentials, iron availability, carbon sources, quorum sensing
molecules, temperature, small organic compounds, and growth phase.
(Finlay and Falkow, 1997; Wilson et al., 2002). In only a few cases is the
precise mechanism of signal perception and transduction known. These
signals may turn on or off the various virulence functions at any level of
regulation, i.e. transcriptional, post-transcriptional, translational and
post-translational. Transcriptional regulation appears to be the most
frequently used because it is the most energetically cost-effective for the
microbe. It is also the more commonly studied because is easier to identify by genetics means. A great variety of regulatory circuits have
been described, from systems where one master regulator is controlling
many virulence genes to pathways where many regulators are involved in
controlling one or a few virulence functions. The biochemical cross-talk
that develops during pathogenic interactions implies that many bacterial
genes are required at various stages of the infection and multi-component
regulatory cascades are needed to integrate the many signals to which
they respond.
Various classes of regulators have been described in bacterial pathogens
(Table 1.1). Among these we find two-component signal transduction
systems, alternative sigma factors (σ 54, σ 32, σ F, σ 28 and σ E-like ECF
sigmas), quorum sensing transcriptional regulators, AraC-like activators,
LysR-type regulators (e.g. Salmonella SpvR, Agrobacterium OccR, and
64 Rhizobium NodD), other activator/repressors directly controlled by small
molecules or ions (e.g. CAP and Fur), and global regulators affecting the
local DNA topology (IHF and H-NS). The remainder of this section will
focus on the generalities of two-component systems.
1.3.2 Two-component systems
Two-component systems (TCS) were first described in the mid-1980s
(Nixon et al., 1986; Ninfa, 1986; Hess, 1988). Since then, hundreds of
TCSs have been described in eubacteria and archea, and to a lesser extent in yeasts, filamentous fungi, and plants (Stock et al., 2000). In bacterial species for which genomic information is available, there are from none
(Mycoplasma genitalium) to 80 (Synechocystis spp.) TCSs. In E. coli, there are at least 30 TCSs (Mizuno, 1997). Many reviews (Stock, 1989;
Bourret et al., 1989; Parkinson, 1992; Stock et al., 1994; Hoch, 2000;
Stock et al., 2000) and books (Hoch et al., 1995) are available on the topic. Here I will briefly introduce some of the structural and functional properties of bacterial TCSs.
TCSs couple signal perception to transcriptional activation of genes (or post-translational activation of enzymes and molecular machines) as a way for bacteria to adapt to diverse niches. TCSs are pairs of proteins with a modular structure (Fig. 1.9). In the simplest case, a sensory histidine kinase (HK) protein, with an N-terminal, sensing (input) domain and a C-terminal conserved kinase core (transmitter) domain, interacts
65 with a response regulator protein (RR), composed of a conserved N-
terminal, regulatory (receiver) and a variable C-terminal effector (output) domain. The RR protein is usually a transcription factor, but there are
exceptions to this general rule. The common theme is that activation of
the RR triggers specific downstream responses. Transduction of stimuli
occurs via phosphorylation reactions involving conserved amino acid
residues in the transmitter and receiver domains. The chemistry of TCSs
includes three phosphotransfer reactions involving His and Asp residues
(Hoch and Silhavy, 1995; Stock et al., 2000):
Autophosphorylation: HK-His + ATP ⇔ HK-His~P + ADP
Phosphotransfer: HK-His~P + RR-Asp ⇔ HK-His + RR-Asp~P
Dephosphorylation: RR-Asp~P + H2 O ⇔ RR-Asp+ Pi
All of these reactions require divalent cations (Mg2+ in particular) for
proper geometry at the catalytic site. HKs catalyze the formation of
phosphoramidate (N~P) bonds, which have a highly negative free energy
of hydrolysis. N~P bonds are acid labile and alkali stable. HKs are
preferentially unphosphorylated due to the thermodynamic properties of
the N~P bond and the normally low ATP/ADP ratio in the cell (Stock et
al., 2000). Perception of the stimuli by the input domain of the HK
changes this equilibrium and the direction of the phosphoryl group flow.
RRs receive the phosphoryl group from their cognate HK forming a high-
energy acyl phosphate bond. Asp~P bonds are acid and alkali labile. The
energy of the Asp~P residues in the context of the protein structure
66 increases the free energy of hydrolysis and this is thought to drive the
long-range conformational changes used by RRs for activation of their
function (Stock et al., 2000). The half-life of the Asp~P moiety of
bacterial RRs in vitro ranges from a few seconds to several hours. It is
often further reduced by the intrinsic autophosphatase activity of the RR
protein, by the presence of phosphatase activity in the cognate HK, or by
the action of cellular regulatory phosphatases (Hoch et al., 1995) (Fig.
1.10).
In addition to the prototypical His-to-Asp phosphotransfer, more complex pathways may be employed to transfer Pi from ATP to the
response regulator. In particular, so called hybrid sensor kinases
(described below) and multicomponent phosphorelay systems use a three-
step His-to-Asp-to-His-to-Asp cascade, which implies the existence of
five distinct reactions (autophosphorylation, three phosphotransfer
reactions and a dephosphorylation step) (Stock et al., 2000). In
phosphorelay systems, the His- and Asp-containing signaling modules are
on separate proteins, instead of being covalently linked. This kind of
signaling molecule is exemplified by the KinA/Spo0F/Spo0B/Spo0A
sporulation cascade of Bacillus subtilis and they are very common in
eukaryotic TCS transduction systems (Stock et al., 2000).
67
1.3.2.1 Response regulators
Response regulators are the “terminal effectors” of the adaptive
response (Stock et al., 2000). They can catalyze the phosphotransfer independently of the HK, as shown by the ability of acetyl phosphate, carbamoyl phosphate, imidazole phosphate and phosphoramidate to function as phosphodonors to RRs (Lukat et al., 1992). Most RRs are transcription factors with a DNA-binding domain in the output region.
Some have enzymatic activities associated with the output domain (e.g. E.
coli CheB methylesterase) and others have no output domain, but are able
to interact with other proteins modulating their activity (e.g. E. coli CheY
interacting with FliM in the flagellar motor) (Fig. 1.9).
The receiver domain is conserved in all RRs. The crystal structures of
several Rrs, such as CheY, NarL and OmpR, show a doubly-wound (α/β) 5 fold. Conserved amino acid residues are located around the catalytic site: a phosphorylated Asp is present at the end of β-strand 3, two Asp residues are in loop 1, a Lys is in loop 5, and a Thr is in loop 4. (In CheY, these would correspond to D12, D13, D57, T87 and K109.) The three conserved
Asp side chains form a catalytic triad coordinating Mg2+ and they are
involved in the phosphorylation and dephosphorylation reactions. Usually,
the most C-terminal Asp residue forms the acyl~P bond. Invariant Thr and
Lys residues also appear to be involved triggering in phosphorylation-
induced conformational changes (Stock et al., 2000).
68 The output domains of RRs with DNA-binding activities are classified
into three different families. OmpR-like regulators form the so-called
“winged-helix” subfamily, the largest group of RRs, based on the analysis
of all the sequenced bacterial genomes (Hoch and Varughese, 2001). This
subfamily is characterized by a DNA-binding fold containing a
recognition helix that binds the major groove of DNA and two flanking
loops (the “wings”) that recognize the minor groove. The mode of action
of different RRs varies, with some recognizing and activating different
portions of the RNA polymerase holoenzyme/promoter complex (e.g.
αCTD or σ 70). In the case of PhoB, phosphorylation promotes
dimerization of the protein (McCleary, 1996; Fiedler, 1996). Direct DNA
repeats are often the binding site for this class of RR and these proteins
bind DNA as asymmetrical dimers (Rampersaud et al., 1989; Harlocker et
al., 1995; Harrison-McMonagle et al., 1999). FixJ-like regulators have
helix-turn-helix DNA-binding motifs in their output domains similar to
the LuxR/GerR-subfamily of transcription factors (Henikoff et al., 1990).
For some members of this class (e.g. UhpA and NarL/NarP),
phosphorylation has been shown to promote symmetrical dimerization and
binding to inverted repeat DNA sequences (Stewart, 1993; Olekhnovich and Kadner, 2002), but direct repeats flanking symmetrical dyads are also targets for cooperative binding to large promoter regions (Li et al., 1994).
Finally, the third subfamily includes NtrC-like RRs, which have output regions composed of an ATPase domain and a helix-turn-helix DNA
69 binding domain (Hoch and Silhavy, 1995). The mechanism of activation
for NtrC has been extensively studied (Keener and Kustu, 1988; Su et al.,
1990; Weiss et al., 1991; Klose et al., 1993; Porter et al., 1993; Klose et
al., 1994; Wedel and Kustu, 1995; Wyman et al., 1997; Pelton et al.,
1999; Rombel et al., 1999). Unphosphorylated NtrC dimers can bind DNA
and the major dimerization domain is located in the C-terminal region.
Phosphorylation, by the NrtB kinase or small phosphodonors, promotes
dimer-oligomerization to larger complexes. Mutations at the conserved
D54, which is the site of phosphorylation, to glutamate mimic the D~P
residue. Finally, NtrC oligomers tethered to the enhancer sites stimulate
ATP hydrolysis, which supplies free energy for the RNA polymerase
holoenzyme/σ 54-promoter isomerization from a closed to an open
complex, activating transcription.
1.3.2.2 Sensor histidine kinases
Sensor HKs are modular proteins located either in the cytoplasm or,
more commonly, transmembrane (Fig. 1.9). They perceive
environmental/cellular signal(s) and transduce them to a cognate response
regulator located in the cytoplasm. HKs range in size from 40 to 200 kDa
and contain from two to six distinguishable domains. In the simplest case, the input domain of the HK controls the phosphorylation events occurring at the kinase core domain by altering its rate of autophosphorylation. In some cases, separate His phosphotransfer (HPt) domains are present.
70 These are able to undergo phosphorylation, but unable to show kinase or phosphatase activities, when separated from the remaining domains of the
HK. In transmembrane sensor kinases, there are often linker domains of
40 to 120 amino acids connecting the sensing/transmembrane domains to the cytoplasmic transmitter and these are important for signal propagation
(HAMP; Appleman et al., 2003; Appleman and Stewart, 2003).
Autophosphorylation is a bimolecular reaction involving cross- phosphorylation of HK dimers at conserved His residues located in their kinase core domains (Stock et al., 2000). In some cases, the HK core also catalyzes the dephosphorylation reaction (e.g. NtrC and FixL), which accelerates the return to the RR “ground” state.
The kinase core of prototypical HKs is about 350 amino acids long and is composed of a dimerization domain and an ATP/ADP-binding phosphotransfer domain or catalytic domain (Stock et al., 2000). The kinase core contains five recognizable amino acid motifs (Hoch and
Silhavy, 1995): the phosphorylation site (H box) and the N, D, F and G boxes in the nucleotide binding site. The dimerization domains form antiparallel four-helix bundles and each usually includes the H box in helix 1. In a few kinases, such as CheA, the H box is separated from the dimerization domain and located in a more N-terminal His- phosphotransfer domain (HPt) (Fig. 1.9). The catalytic domain contains the N-D-F-G boxes and binds ATP/ADP at a highly flexible cleft. Most structural information is based on the E. coli CheA and EnvZ protein
71 fragments derived from X-ray diffraction crystal structures and/or NMR
solution structures, but a complete structural analysis for a full length HK
is not available yet.
The kinase core interacts with the receiver domain in the RR during the
phosphotransfer step. How these two domains interact has been partially
revealed by cocrystalization of the Spo0F response regulator with the
cognate Spo0B phosphotransferase (Zapf et al., 2000). The interaction surface involves residues on the loops connecting the α-helices and β- strands of the RR and the α1 and α2’helices in the dimerization/H box of
the HK (Hoch and Varughese, 2001). Invariant residues are involved in
the catalytic process. Such residues are brought into the proper geometry
upon the interaction of conserved anchor residues and variable
recognition residues at the interaction surface. The variable recognition
residues are responsible for preventing heterologous interactions among
non-cognate HK-RR pairs, thereby ensuring specificity in signal
propagation (Zapf et al., 2000).
Based on the domain organization, at least two major classes of HK
have been recognized. Orthodox histidine kinases are either soluble or,
more commonly, transmembrane proteins composed simply of an input
sensory domain and a cytoplasmic transmitter domain (Fig. 1.10). Most
HKs fall into this class. Hybrid sensor kinases are composed of a set of
additional RR-like receiver and HPt domains at the C-terminus of an
orthodox HK structure. This domain architecture allows for an
72 intramolecular His-to-Asp-to-His phosphotransfer cascade. This general
organization is found in several hybrid kinases such as E. coli ArcB,
Salmonella spp. BarA, and Bordetella BvgS, but in a few cases either the
HPt domain is missing (E. coli RcsC, Mixococcus spp. FrzE) or
duplicated/triplicated receiver domains are present, with (Vibrio cholerae
NP_230993) or without (X. campestris NP_642807; P. putida NP_744503)
an HPt domain (CDART analysis at the NCBI database).
Input sensory domains are essential components of HKs but in general
they are poorly characterized. As mentioned above, in the majority of the
cases, the nature of the specific signals perceived at the molecular level
are not known. Moreover, the mechanisms by which the sensory domain
controls the autophosphorylation rate at the transmitter domain are also
not well understood. In some cases, deletion of the input domain releases
the autokinase activity of the sensor protein (Monson et al., 1992;
Scholten and Tommassen, 1993; Kramer and Weiss, 1999), whereas in
other systems reduces the autophosphorylation activity (KinA; Wang et
al., 2001) or specifically increases the phosphatase activity (E. coli
KdpD; Jung and Altendorf, 1998). Input domains generally show little
sequence conservation, even if a few families of signaling motifs have
been recognized. The PAS (Per-Arnt-Sim) domain superfamily (Pfam
00989) is perhaps the best characterized (Zhulin, 1997; Taylor, 1999;).
PAS domains have been found in over 1100 proteins encoded by both prokaryotic and eukaryotic genomes. PAS domains are cytoplasmic, small
73 (100-120 amino acids) protein motifs that fold into a conserved PAS core,
containing a β-scaffold and a helical connector (See Chapter 5 for more
details on their structure). The structural elements of the mixed α/β fold
are designated A , B , C , D , E , F , G , H , and I and a standardized
numbering system has been proposed to facilitate comparison of the
various PAS motifs (Amezcua et al., 2002). PAS domains are commonly
involved in protein/protein interactions and intermolecular association of
macromolecules (Amezcua et al., 2002). They also function as sensory
modules. Various cofactors, which determine signal specificity, are
accommodated within the hydrophobic PAS core. PAS domains have been
shown to bind many different types of cofactors, such as heme, to sense
O2 (FixL), 4-hydroxycinnamic acid to sense blue light (PYP), FAD, to
sense redox levels (NifL), FMN, to sense ble light (plant phototropins)
and ATP, to sense the energy charge (KinA) (Stephenson and Hoch,
2001). In bacteria, PAS motifs are exclusively found in TCSs, as single,
tandem or multiple domains (up to six). The presence of multiple PAS
motifs in a sensor raises the interesting possibility of multiple ligands
being perceived by a single protein. Due to their importance both as
universal sensory modules and as key components of important bacterial
virulence regulatory systems, PAS domains are being characterized in
great detail using biophysical methods. Intramolecular conformational
changes upon ligand binding have been recently observed in solution
using NMR (Amezcua et al., 2002). In particular the FG loop has been
74 shown to be critically important for the inhibition of the kinase core activity in the absence of a ligand in the PAS core pocket. Binding of hydrophobic ligands would trigger conformational changes that propagate to the Fα helix, the FG loop and the Hβ strand, thereby relieving the
inhibition of the kinase core (Amezcua et al., 2002).
Other families of motifs are indentifiable in input sensory domains of
TCS kinases (reviewed in more detail by Galperin et al., 2001). For some, the ligand is known or hypothesized. These motifs include the FliY-like periplasmic binding protein (PBPb) domain (~220 aa; Pfam 00497), often involved in amino acid binding, the Cache domain (~120 aa; Pfam02743), the MHYT module (~200 aa; COG3300), the GAF sensor domain (~150 aa; Pfam 01590), similar to a PAS fold but binding cGMP, the HAMP linker domain (~50 aa; Pfam00672) and the CBS domains (Pfam 00517).
1.4. SIGNIFICANCE, RATIONALE AND OBJECTIVES
1.4.1 Significance
Stewart’s wilt is an economically important disease of sweet corn in the
eastern part of the U.S. “Corn Belt”, especially after mild winters. It can
cause up to 100% yield losses in the field. Virtually no direct control
measures exist and disease management is mainly based on pesticide
treatments to control the insect vector and the use of tolerant sweet corn
cultivars. P. stewartii is a quarantine pathogen for more than 100
countries importing U.S. hybrid corn seed and frequent commercial
75 controversies arise between the U.S.A. and the European Community or
Far East countries on issues related to the presence of P. stewartii in corn
seed stocks. Understanding the virulence mechanism of P. stewartii at a
molecular level will eventually open new avenues for the rational
management of Stewart’s wilt and perhaps for the elimination of
transmission of the pathogen via seed.
P. stewartii-corn pathosystem is a good model for studying hrp type III
secretion apparatuses and their regulation because: (a) P. stewartii is a
good endophyte that can colonize a wide range of grasses without causing
symptoms in many of them, it causes disease in a few hosts, including
cultivated corn, and it multiplies in the intestinal tract of insect vectors.
In contrast to P. agglomerans, P. stewartii is not found in environmental
samples or on the surfaces of plants because it survives poorly under
water stress and in sunlight. (b) P. stewartii uses two of the major
virulence mechanisms of phytobacteria, i.e the production of EPS to plug
water-conducting xylem vessels and the production of effector proteins to
cause watersoaking of parenchymatic tissues. This offers the possibility to
study the coregulation of two important pathogenicity mechanisms (cps
and hrp genes). (c) The master regulators of hrp genes are conserved in
all erwinias. Therefore, we hope that any information gained from P.
stewartii may be applicable to all other members of this group. (d) Most
genetic tools developed for E. coli genetics are applicable to P. stewartii.
(e) The P. stewartii genome is being sequenced at the Baylor College of
76 Medicine and the University of Wisconsin at Madison and a draft sequence should be available at the begining of 2004. This will greatly facilitate functional analysis of pathogencity genes in this species and the study of their regulatory networks. (f) Corn is an ideal experimental host because it is easy to grow and and most of the Stewart’s wilt symptoms can be observed on young seedlings. (g) Corn is also one of the best genetically characterized crops; detailed genetic maps, vast EST libraries and numerous molecular reagents, including BAC and yeast two-hybrid libraries and transposon mutant collections, are publicly available.
1.4.2 Rationale
hrp genes are essential for the pathogenicity of P. stewartii as they are for many other necrogenic phytobacteria. Several observations make the study of hrp regulation particularly attractive in P. stewartii.
In many non-host plants, P. stewartii often remains latent without eliciting apparent host defense responses even at high inoculum doses.
For example, P. stewartii cells infiltrated at high cell density into tobacco will not cause hypersensitive plant cell death unless the bacteria are pre- grown in minimal media that mimics the metabolic signals that activate hrp/hrc type III secretion genes in planta. Many plants, such as tobacco and various grasses, are apparently unable to provide adequate signals to alert P. stewartii that it is in a potential host. Understanding the molecular mechanisms by which hosts and the environment control the
77 expression of virulence determinants may open the way to new strategies
for disease control, for instance by interfering with the sensory system of
the pathogen before an R-Avr interaction is even established.
Metabolic control of virulence gene expression plays an important role
in group I hrp genes but little is known about the earliest steps of signal
sensing. The hypothesis put forward in this dissertation is that the HrpX
PAS-kinase and the HrpY response regulator form a signal transduction
system that senses nutritional/energy stresses. The discovery of potential
HrpX/HrpY orthologs in the unfinished genome sequences of X. campestris and P. syringae DC3000 further suggests that control by starvation signals may be common to all group I and II hrp systems and perhaps to other pathogens. Once the genome of P. stewartii is sequenced, the regulatory networks that cross-talk with the HrpX/HrpY system and other downstream master regulators may be dissected. The molecular characterization of the regulatory cascade involving HrpX/HrpY, HrpS and HrpL will provide a solid framework for future studies at the genome- wide level.
Quorum sensing regulation of P. stewartii capsule polysaccharide production has been well characterized (Von Bodman, et al., 1998).
Mutants unable to produce OHHL produce very little EPS and are severely affected in their ability to cause wilting and watersoaking in corn seedlings. While the original report on esaI mutants focused on EPS production, the reduction in watersoaking due to this mutation suggested
78 that quorum sensing may also affect hrp gene expression, because cps mutants are still able to form lesions. This may imply a novel modality of host-pathogen interaction in the early phases of pathogenesis. At the same time it opens new avenues for controlling Stewart’s wilt by targeting the quorum sensing signaling mechanism.
1.4.2 Objectives
This dissertation deals with several aspects of the signal transduction cascade that activates and/or represses hrp regulatory genes in P. stewartii. In particular, I focused primarily on the regulation of hrpS by the HrpX/HrpY two component and the EsaR/EsaI quorum sensing systems and secondarily on the regulation of hrpL by HrpY. HrpS and
HrpL are two key transcription factors that control the activation of all known hrp/hrc/wts genes in P. stewartii. The HrpX/HrpY system is predicted to mediate a typical adaptive response to environmental signals and the EsaR/EsaI quorum sensing system mediates cell-to-cell communications.
The major objectives of my research were:
1. To genetically dissect the regulatory cascade controlling
hrp/hrc/wts gene expression in P. stewartii, which involves the
HrpX sensor kinase, the HrpY response regulator, the HrpS
enhancer-binding protein and the HrpL alternative sigma factor
(Chapter 2; published in Merighi et al., 2003).
79 2. To analyze at the genetic and biochemical levels a novel
autoregulatory loop triggered by HrpS (Chapter 3).
3. To analyze the biochemical properties of the response regulator
HrpY and its novel dual role as activator of hrpS and a repressor of
hrpL (Chapter 4).
4. To genetically dissect the sensory domains of HrpX and establish
reagents for the future study of its biochemical functions (Chapter
5).
5. To define the classes of signals perceived by the various
components of the regulatory cascade controlling hrp genes
(Chapter 5).
6. To determine the role of AHL-based quorum sensing in modulating
hrp gene expression in P. stewartii (Chapter 6).
While the data presented in Chapters 2, 3 and 6 can be considered completed studies, Chapter 4 and especially Chapter 5 contain much preliminary data that will be valuable for future studies.
80
Organism AraC-like Two- Alternative Histone- Other proteins component sigma like factors systems factors proteins Yersinia spp. VirF=LcrF YmoA S. typhimurium InvF HilA SPI-1 BarA/SirA PhoQ/PhoP S. typhimurium SpiR/SsrA, SPI-2 EnvZ/OmpR
E. amylovora HrpS HrpL HrpX/HrpY σ54? P. syringae pv. HrpR, HrpS HrpL syringae σ54 R. solanacearum HrpB HrpG PhrI PhrR PhrJ PhrA X. campestris HrpXv HrpG
E. carotovora HrpS HrpL RsmA/B, GacS/GacA σ54 IHF
Table 1.1 Regulators of type III secretion systems.
81 . y endemic; Black dots indicate interceptions and/or interceptions indicate dots Black endemic; ort of the disease in the countr the in disease of the ort p
oradic outbreaks. The dates refer to the first re p
. Figure 1.1. Worldwide distribution of Stewart’s wilt. is Red (grey) dots indicates areas were the disease s
82 6
4
5 7 9
3
1 8 2
2 mm
1
Figure 1.2. Stewart’s wilt disease cycle.
Bacteria overwinter in the flea’s beetles (2) alimentary tract or in corn seeds (1).
Survival in asymptomatic hosts may also occur. Corn seedlings can be infected via seed transmission, leading to early stunting, wilting and leaf streaking (5). Beetle infestations transmit the disease through feeding wounds on the leaves (4). The vascular bundles (6) and the intercellular spaces (9) are colonized, watersoaking occurs and dense population can be seen in xylem vessels (7). Bacteria can be seen streaming out of xylem vessels in transversal sections of infected leaves (8). Leaves show elongated, lens-shaped lesions that become necrotic. Secondary infections can be started by contaminated beetles.
83
Apo-EsaR OHHL EsaR active
inactive
Acyl-carrier protein SAM EsaI
EPS
esa I esaR
esaR-Box
Figure 1.3. The quorum sensing regulatory circuit controlling EPS synthesis in P.
stewartii.
The LuxR-homolog EsaR receptor-regulator binds target DNA elements as an
apoprotein. EsaI is a synthase catalyzing the formation of OHHL from SAM and acyl-
carrier protein. Increased OHHL concentration at high bacterial cell densities titrates
out EsaR. The OHHL-conjugated EsaR has much lower affinity for the target promoter
elements. The model is analogous to the lacI/lacZ operon model. OHHL: oxo- hexanoyl-homoserine lactone; SAM: S-adenosyl-methionine.
84 N C N C N C N C
OM PulD
N C Periplasm N C N
IM Sec
N C N C N C N C Chaperones Chaperones Type I II III IV V
Fig. 1.4. Bacterial secretion systems.
Five major classes of bacterial secretion systems are recognized. Type I systems are members of the ABC transporter superfamily. They recognize substrates via C- terminal signals ans translocated them in a single step. Type II secretion systems utilize the inner membrane Sec apparatus (general secretory pathway) to recognize N- terminal cleavable signal pepetides in their substrates. Secretion involve a periplasmic intermediate. Translocation through the outer membrane requires a multimeric secretin-like complex. Type III secretion systems secrete and translocate proteins recognized via various targeting mechanisms (chaperones, 5’-UTR-mRNA and non-cleavable signal peptides). Type III secretion systems are required for host- bacteria symbiotic/pathogenic interactions of many gram-negative species. Type IV systems vectorially secrete proteins or protein/DNA complexes. Type V systems are the so-called autotransporters. Substrates traveling this system cross the inner membrane via the sec pathway. IM inner membrane; OM, outer membrane (Adapted from Cossart et al., 1999),
.
85 YopB
YscD
20 Å 1200 Å 70 Å DISTAL ROD L RING
o.m. o.m.
YscC P. 50 Å PROXIMAL P RING 200 Å BASAL ROD BODY MS RING YscJ*
i.m. i.m.
C RING YscQ
YscN Flagellum YscU Specific secretion YscT YscR apparatus YscS LcvD
Hrp Pilus
o.m.
HrcC P.
HrcJ*
i.m.
HrcQ
HrcN o.m.= outer membrane i.m. = inner membrane p. = peptidoglycan layer HrcU HrcT HrcS HrcR HrcV
Figure 1.5. Schematic representation of animal and plant pathogen type III secretion apparatuses and of the flagellar basal body. Adapted from Cossart et al., (1999) and He, (1998).
86
s are regulatory genes, blue boxes are conserved genes, blue s are regulatory al and plant pathogen type III secretion genes. secretion genes, white boxes are other secretion genes secretion boxes are other genes, white secretion Figure 1.6. Transcriptional rganization of anim red boxe boxes Black proteins, are genes for secreted
87 Yersinia spp. 37ºC yop/ysc/ lcrGV/sycD
virF (escape macrophage endocytosis) LcrQ 26ºC
YmoA=H-NS
S. enterica Typhimurium (SPI-1) ? Supercoiling ~P +
barA sirA hilA invF sip/ssp inv-spa-prg (epithelium invasion)
Low pH High Fe3+ 2+ Low Mg RcsA/B/C
~P
phoQ phoP pmrD pmrB pmrA ugd (LPS modification)
Low pH ? Low osmolarity
~P (SPI-2) ~P
ompR envZ ssrBA spi- ssa- sse
(macrophage survival & proliferation)
Figure 1.7. Regulatory circuits controlling type III secretion system transcription in animal pathogens.
Red (dark grey): sensor kinases; blue (light grey): response regulators; white: AraC- like activators; black arrows: secretion genes.
88
Figure 1.8. Phytobacterial Hrp/type III secretion system regulatatory circuits.
Red (dark grey): sensor kinases; blue (light grey): response regulators; white: AraC- like activators; yellow (stripes), alternative sigma factors; black arrows: secretion genes; i.m. inner membrane; o.m., outer membrane.
89
P. syringae Lon RpoN IHF ENVIRONMENTAL SIGNALS Hrp Box +/ hrpR hrpS hrpL hrp/hrc ? ? HrpA HrpV
E. amylovora ?
Hrp Box
hrp/hrc hrpX hrpY hrpS hrpL
P. carotovorum Constitutive RpoN IHF ? Hrp Box
hrp/hrc hrpX hrpY hrpS hrpL GacA/S RsmA OHHL
X. campestris ?
(PIP) hrcC UNKNOWN hrpG hrpX hrp/hrc
(Figure 1.8 continued)
90 Figure 1.8 (continued)
Metabolic signals R. solanacearum PLANT CELL CONTACT PhrI/PhrR
PrhA (TonB-like) hrp/hrc i.m. phrJ hrpG hrpB on the o.m.
91
Figure 1.9. Modular structure of two-component systems.
Two-component sensor kinases and response regulators are formed by modular domains performing the discrete tasks of signal perception, transmission, reception and output response. Sensor kinases can be composed of more that two modules allowing for intramolecular phosphotransfer cascades. In class I kinases the H-box containing the phosphrylatable Histidine residue is located nearby the glycine-rich
ATP binding domain; in class two kinases the Hrp domain is physically separated from the transmitter domain. Pfam and COG protein domain databases are available at www.sanger.ac.uk/software/ Pfam and www.ncbi.nlm.nih.gov/cog. (Figure adapted from Swanson et al., 1994).
92
D CheY, Spo0F
OmpR, and FixJ classes D (e.g. HrpY, NarL, UhpA, SirA) Receiver D NtrC class D (enhancer-binding ) ~120 aa D Methyl-transferases; GGDEF, EAL, HD-GYP regulators Response regulators
~ H D H ArcB, BarA
Hybrid kinases
~ H D RcsC
Class I H NarX
H EnvZ H Kinase core H Rm FixL ~250-350 aa
NtrB, KinA, DegS, HrpX H Bj FixL
H CheA Class II
Orthodox kinases Figure 1.9 (continued)
93
Figure 1.9 (continued)
Legend:
D Receiver (Pfam00072)
σ54-dependent ATPase (Pfam00158)
GGBEF (Pfam01590), EAL (Pfam00990), HD-GYP (COG2206)
DNA binding : GerR-like HTH (Pfam00196), Fis-like HTH (Pfam02954.9); winged
helix (COG0745)
H HisKA or HATPase (Pfam00512 and Pfam02518) H Hpt (Pfam01627) Periplasmic domain (e.g. Pfam00497, Pfam02743, COG3300; etc.)
Cytoplasmic input domain (often with PAS/PAC domains; COG2202)
H-kinase dimerization domain (Pfam 2895.9)
HAMP linker (Pfam00672)
Rm, Rhizobium meliloti; Bj, Bradyrhizobium japonicum; H, Histidine; D, aspartate; Hpt, histidine phosphotransfer; HTH, helix-turn-helix
94
P P
D PhoP H NDFG ATP PhoQ ADP Pi H2O (Orthodox SK) P Class I D CheY P
H NGNDFGFG P 1 2 CheA D CheB (Orthodox SK) Class II
P P P P
H NDFG D H D BvgA NG1FG2 BvgS (hybrid SK) H PPP P
H NDFG D H D RcsB NG1FG2 RcsC YojN (hybrid SK) P
H NDFG P P P KinA D H D Spo0A P Spo0F Spo0B H NDFG (Phosphorelay) KinB
Figure 1.10. Phosphorylation pathways in two-component systems (Adapted from
Stock et al, 2001).
95 CHAPTER 2
THE HRPX/HRPY TWO-COMPONENT SYSTEM ACTIVATES HRPS
EXPRESSION, FIRST STEP IN THE REGULATORY CASCADE
CONTROLLING THE HRP REGULON IN PANTOEA STEWARTII
SUBSP. STEWARTII
2.1 INTRODUCTION
Pantoea stewartii subsp. stewartii (Basonym Erwinia stewartii) causes
Stewart’s bacterial wilt and leaf blight of sweet corn and maize (Zea mays
L.). Stewart’s wilt is the most serious bacterial disease of sweet corn in the north-central and eastern United States. The pathogen is transmitted by the corn beetle (Chaetocnema pulicaria) through feeding wounds
(Claflin, 1999). Bacteria grow in the xylem vessels of the host plant causing wilt and in the intercellular spaces of the leaves causing “water- soaked” lesions. The primary virulence factor responsible for wilting is the extracellular polysaccharide (EPS; Dolph et al., 1988) and the genes required for its production and regulation have been characterized
(Torres-Cabassa et al., 1987; Dolph et al., 1988; Coplin and Majerczak
96 1990; Poetter and Coplin, 1991). Water-soaking (Wts) is the result of cell
membrane damage and accumulation of fluids in the apoplastic spaces of
the leaf parenchyma.
A cluster of hrp genes (Alfano and Collmer, 1997) is required by many
gram-negative plant-pathogenic bacteria for elicitation of the
hypersensitive response (HR; Klement 1982) in nonhost plants and for
pathogenicity in a compatible host. hrp genes encode a type III secretion
system that is conserved among plant and animal bacterial pathogens
(Galan and Collmer 1999). They also encode several regulatory proteins
and the secreted effector proteins that determine interactions with
susceptible and resistant hosts. P. stewartii has a typical hrp cluster that
spans over 28-kb and is required for HR elicitation in tobacco and disease
in corn (Coplin et al., 1986; Coplin et al., 1992; Ahmad et al., 2001;
Frederick et al., 2001). In contrast to more necrogenic pathogens, such as
Pseudomonas syringae and Erwinia amylovora, P. stewartii is only able to elicit an HR in tobacco leaves when the bacteria are pre-grown in a Hrp- inducing, minimal medium or hrpS is constitutively expressed (Ahmad et al., 2001). Thus unlike sweet corn and maize, tobacco does not provide all of the signals necessary to induce hrp gene expression in this species.
In gram-negative plant pathogenic bacteria, there are two lineages of hrp genes that differ significantly in their mode of regulation (Alfano and
Collmer 1997). In general, expression of hrp genes is environmentally
regulated. Both plant factors (Schulte and Bonas 1992; Aldon et al., 2000)
97 and chemical/metabolic stimuli mimicking the plant apoplast (Rahme et
al., 1992; Wei et al., 1992; Xiao et al., 1992) have been shown to activate
their expression, but the mechanism by which these signals are perceived
is still unknown. Group I hrp genes in P. syringae, Erwinia spp.,
Pectobacterium spp. and Pantoea spp. are regulated by HrpS and the
alternative sigma factor HrpL (Frederick et al., 1993; Xiao et al., 1994;
Xiao and Hutcheson 1994; Wei and Beer 1995). Expression of hrpL
requires the alternate sigma factor σ 54 in addition to the NtrC-like
transcriptional activator HrpS (Frederick et al.,, 1993; Hendrickson et al.,
2000a). In P. syringae, HrpR, which is homologous to HrpS, is also
required for full activation of the hrpL promoter and may form
heteromeric complexes with HrpS (Hutcheson et al., 2001). In E.
amylovora, Wei et al., (2000b) described a two-component signal transduction system, HrpX/HrpY, which is orthologous to the one described in this work. They proposed that both HrpS and HrpY regulate
hrpL and their effects are additive. In contrast, Group II hrp genes in
Xanthomonas and Ralstonia spp. are regulated by AraC-like
transcriptional activators. In Ralstonia solanacearum, a complex
regulatory cascade transduces a signal arising from contact with the plant
cell wall and activates hrp gene expression (Aldon et al., 2000; Brito et
al., 2002).
In this report, we describe the sequences and function of genes included
in P. stewartii hrp complementation groups V-VII, which were previously
98 identified as regulatory loci (Frederick et al., 2001). hrpS (formerly designated wtsA) was characterized by Frederick et al., (1993) and its sequence is reanalyzed here in the context of new data. Using mutational and epistasis analysis, we determined the Hrp phenotype of individual regulatory genes and the order of the steps in the regulatory cascade. The results of this study support a model in which phosphorylated HrpY activates hrpS in response to environmental stimuli. This is different from the pathway reported for E. amylovora, but similar to that concurrently discovered in Pantoea agglomerans pv. gypsophilae (Nizan-Koren et al.,
2003).
2.2 MATERIALS AND METHODS
2.2.1 Bacterial strains, plasmids, media and growth conditions
E. coli and P. stewartii strains and plasmids used in this study are listed in Table 2.1. Luria-Bertani (LB) broth and agar (Ausubel et al., 1987) were used for strain maintenance and for growth of P. stewartii under
Hrp-repressing conditions. Fusaric acid-containing tetracycline sensitive selection (TSS) medium was prepared as described (Metcalf et al., 1996).
To induce hrp genes in P. stewartii strains, overnight LB broth cultures were washed twice in inducing medium (IM; Frederick et al., 2001), inoculated into 2 ml of the same medium at OD600nm = 0.05, grown for 12-
16 h and adjusted to final OD600nm ca. 0.5. In the pH experiment, HEPES and Tris-HCl buffers were used for the neutral and basic pH ranges,
99 respectively, in place of MES. Liquid cultures were grown in flasks or tubes shaken at 200 rpm at 37°C for E. coli or 29°C for P. stewartii.
When appropriate, antibiotics were supplied at the following
concentrations: ampicillin, 200 µg ml-1; kanamycin, 50 µg ml-1; and tetracycline, 20 µg ml-1 (or 15 µg ml-1 for selection of single copy Tetr cointegrates).
2.2.2 HR and pathogenicity tests
To test for HR elicitation on tobacco (Nicotiana tabacum L., cv. “White
Burley”) plants, P. stewartii strains were grown overnight in IM liquid
9 medium, washed and resuspended in water at OD600nm = 0.52 (ca. 1x10
CFU ml-1). Cells were infiltrated into tobacco leaves using four replicates
per strain as previously described (Frederick et al., 2001) and necrosis
was rated 24-48 h after infiltration. Pathogenicity tests were performed by
inoculating the whorls of 5-day-old sweet corn seedlings (Zea mays var.
saccharata, cv. “Seneca Horizon”) as previously described (Coplin et al.,
1986). The plants were held in growth chambers at 29°C (photoperiod 16
h, 15000 lux, relative humidity 99%). After 3 days, disease severity was
rated using a 0 to 3 scale (0= no symptoms, 1=scattered small lesions, 2 = numerous lesions, and 3 = extensive lesions that remained water-soaked with ooze forming on leaf surfaces).
100
2.2.3 General recombinant DNA, genetic and bioinformatic techniques
DNA isolation, agarose gel electrophoresis, restriction enzyme
digestion and ligation were done following standard procedures (Ausubel
et al., 1987). PCRs used either native Taq (for colony-PCR screens and
construct mapping) or cloned Pfu (for molecular cloning) DNA
polymerases (Invitrogen, Carlsbad, CA, U.S.A and Clontech, Palo Alto,
CA, U.S.A., respectively) and standard amplification protocols (Ausubel
et al., 1987). For colony-PCR, we used cells lysed in 50 mM NaOH for 10
min at 100°C as templates. For pBluescript and pRK415 vectors, the
reactions were performed using one primer anchored on either the lacI
vector gene (primer LACF) or the αlacZ gene fragment (primer LACR)
and a second primer conveniently located in the recombinant insert (Table
2.2). Standard deprotected oligonucleotide primers were ordered from
IDT-DNA. Probes for Southern blot hybridizations were labeled with 11-
Dig-dUTP and colorimetric immunodetection was done as recommended by the manufacturer (Roche Biochemicals). Plasmids were introduced into
E. coli and P. stewartii strains by electroporation, chemical transformation (Inoue et al., 1990) or mobilized by triparental mating using pRK2013::Tn7 as a helper plasmid (Ditta et al., 1980).
DNA sequencing was performed using cycle sequencing with fluorescent dye terminator chemistry and the ABI 3700 capillary electrophoresis sequencer (PE-Applied Biosystems) at The Ohio State
101 University Plant-Microbe Genomic Facility. Database analyses of
predicted proteins were performed using BLAST/PSI-BLAST (Altschul et al., 1997), whereas conserved domain analysis used the CD-search service
(ver. 1.54; Marchler-Bauer et al., 2002), both available through the
National Center for Biotechnology Information
(http://www.ncbi.nlm.nih.gov). Statistical significance of the matches are reported as E-values. Promoter searches were performed using a neural
network prediction algorithm (NNPP; Reese et al., 1996) and the
IHF/Sigma54 promoter-finder program (Goodrich et al., 1990). RNA
secondary structure predictions were performed at the
www.genebee.msu.su web server. Other sequence manipulation and
analyses were executed using software procedures in the GCG package
Ver. 9 (Oxford Molecular).
2.2.5 Construction of Plac-hrpY, Plac-hrpS and Plac-hrpL plasmids
For complementation and over-expression experiments, the hrpX, hrpY and hrpL ORFs, including their ribosomal binding sites, were amplified by PCR and cloned into high and low copy number cloning vectors such that they were expressed from Plac (Fig. 2.1). E. coli DH10B was used as
a cloning host. In all cases, cosmid pES1044 or plasmid pMM58 were
used as templates for the amplification reactions. PCR products were gel-
purified, digested with appropriate restriction enzymes, and then ligated
into pBluescript KS(+) or pBluescript SK(+) (Stratagene). Clones
102 containing recombinant plasmids were screened by colony-PCR using the
LacF primer and the reverse primer used for the PCR cloning. Individual
genes were subsequently subcloned into the low copy, mobilizable vector,
pRK415 (Keen et al., 1988), such that they were transcribed from Plac.
Vectors and recombinant plasmids are listed in Table 2.1 and primers in
Table 2.2.
hrpY was amplified using primers YFH and YRH. The 675-bp amplicon
was cloned into pBluescript KS(+) to produce pMM46, and then subcloned
into pRK415, to produce pMM52. hrpL, without its σ 54-dependent
promoter, was amplified using primers LFB and LRE. The 613 bp PCR
fragment was cloned into pBluescript SK(+) to produce pMM22 and then
subcloned into pRK415 to produce pMM6. The construction of Plac-hrpS
expression plasmids pRF205 (low copy) and pRF8 (high copy) and the
hrpXY plasmids pRF201 was previously described (Frederick et al., 1993).
Plasmid pMM58, containing the entire regulatory region, was constructed
by ligating the 1.7 kb HindIII insert of pRF8 into pRF201SK digested
with HindIII.
2.2.6 Construction of chromosomal P. stewartii hrp insertion mutants
Construction of Tn5 and Tn3HoHoI mutants with insertions in hrpX,
hrpY, hrpS and hrpL has been previously described (Frederick et al.,
2001). Double mutants containing selected null mutations and lacZ gene fusions were constructed by allele exchange of Tn5-mutagenized cosmids
103 as previously reported (Frederick et al., 2001) by using an incompatible
plasmid, pR751 (Ruvkun and and Ausubel 1981), to force the first cross-
over event. Nonpolar hrpL mutations were constructed by ligating the
aphA-3 cassette from pUC18K (Menard et al., 1993) into hrpL in pRF201.
The resulting plasmid, pDM2560, was used to exchange the hrpL::aphA-3
allele into the P. stewartii chromosome as described above. Several
strategies to construct strains with usable chromosomal hrpL-lacZ
transcriptional fusions were unsuccessful. Therefore, we created a strain
with a single-copy chromosomal hrpL-uidA transcriptional fusion by a
Campbell-type single crossover event. Plasmid pDM2785, which contains
hrpLp cloned into pPL6GUSC, was forced to recombine with the hrpL locus on the chromosome following introduction of an incompatible plasmid, pR751, to produce hrpL-uidA/hrpL+ Tetr Tpr strains. This
plasmid was recombined into wild-type (DC283), hrpY (DM064) and hrpS
(DM758) backgrounds to produce strains DM2831, DM2837 and DM2844,
respectively.
2.2.7 Site-directed mutagenesis of hrpY codon D57 by overlap
extension PCR and construction of missense mutant strains
To evaluate the role of the conserved aspartate residue at position 57 in
HrpY, we performed site-directed mutagenesis by a modification of the
megaprimer overlap extension PCR technique (Aiyar and Leis 1993). D57
was changed by nonconservative substitution to alanine [D57A] or by
104 conservative substitution to asparagine [D57N]. Five primers were
designed: YFH and YRH delimited the hrpY ORF; YR2 was the antisense
primer; and YD57N and YD57A were the sense-orientation mutagenic
primers for each substitution. The mutagenic primers contained missense
base changes at D57 (bold font in Table 2.2) and a near silent restriction
site (NheI and DraI for YD57A and YD57N, respectively). PCRs were
performed using pMM46 as template DNA. The first round of reactions
used primer pairs YFH+YR2 and YRH+YD57A (or YD57N) to produce
350 bp and 500 bp (or 508 bp) overlapping amplicons. Gel purified PCR
products from the two reactions were mixed, annealed and used as templates for a second round of PCR using primers YFH and YRH. PCR products were gel-purified and cloned into pBluescript KS(+).
Recombinant colonies were screened by PCR and by restriction enzyme analysis for the presence of the marker restriction site and the inserts were sequenced. Plasmids pMM57 and pMM92 contain the hrpY[D57A] and hrpY[D57N] alleles, respectively. These inserts were then subcloned into pRK415 to produce plasmids pMM74 and pMM118, respectively. To construct missense hrpY mutants, the hrpY[D57A] and hrpY[D57N] ORFs
were subcloned from pMM57 and pMM92 into suicide vector pLD55
(Metcalf et al., 1996) as XhoI/SstI inserts. The unmarked mutations were
introduced into the P. stewartii genome using the procedure described by
Metcalf et al., (1996) and confirmed by Southern blot hybridization.
105 2.2.9 Construction of P. stewartii strains with in-frame deletions in
hrpX
Two unmarked in-frame deletions in hrpX were constructed in vitro by
PCR. The deletions span amino acids 17 to 414 and 17 to 470 as indicated in the subscripts. To construct ∆hrpX(17-414), DNA fragments
corresponding to the ends of hrpX and its flanking sequences were
amplified by PCR from cosmid pES1044 using primer pairs EdX1+BdX1
and BdX4+XdX3 (Table 2.2). ∆hrpX(17-470) was similarly constructed
using the same primers, except BdX3 was substituted for BdX4. PCR
fragments were ligated into pBluescript KS(+) as EcoRI-BamHI and
BamHI-XbaI contiguous fragments. The deleted alleles were then ligated into pLD55 as XhoI/SstI inserts, transformed into E. coli BW20339
(Metcalf et al., 1996), mobilized to P. stewartii and introduced into the chromosome as described above. Cosmid pDM1296 hrpX+ hrpY::Tn5, which expresses hrpX from its native promoter, restored HR elicitation to these mutants, thereby genetically confirming that these strains are in fact bona fide nonpolar hrpX mutants.
2.2.9 Construction of hrp-promoter reporter plasmids
hrp promoter fragments containing the 5’ regulatory regions of hrpX, hrpS, hrpL, hrpJ and wtsE were either produced by PCR using the primers described in Table 2.1 or isolated as restriction fragments from cosmids pES411 or pES1044. To amplify the hrpL promoter region from -207 to
106 +182 (coordinates refer to the first base of the ORF), primers JFBG and
LRBG were used (Table 2.2). An hrpX promoter fragment containing from
–198 to + 46 bp was produced with primers XFHG and XRBG. The hrpS
promoter region from nucleotides -928 to +384 bp was amplified with primers SFG and SR3BG. To isolate two promoter regions containing hrp
boxes, a 0.9 kb BamHI insert containing PhrpJ and a 1.1 kb HindIII-BamHI
fragment containing PwtsE were purified from plasmid pRF201::Tn5seq118
and cosmid pES411, respectively. The former plasmid has a Tn5seq
insertion at +252 bp in hrpL (R. Frederick and D. Coplin, unpublished).
The DNA fragments, digested with BamHI and/or HindIII as required,
were ligated into the low-copy, promoterless, uidA fusion vector
pL6GUSC (Knoop et al., 1991). hrp-uidA reporter plasmids are listed in
Table 2.1. The same inserts used for hrp-uidA fusions were cloned into
the gfp reporter vectors pPROBE-KT or pPROBEgfp[AAV] (Miller et al.,
2000) to produce plasmids pMM165 hrpS-gfp[AAV], pDM2760 hrpJ-
gfp[AAV], pDM2756 hrpL-gfp[AAV] and pDM2724 hrpX-gfp.
2.2.10 Measurement of enzymatic reporter gene activity
β-glucuronidase (GUS) activity was assayed fluorometrically using 4-
methyl-umbelliferyl-β-D-glucuronide as described by Jefferson (1987),
but the assays were scaled down to fit microtiter plates and analyzed
using a Victor 1420-2 multilabel reader (PE-Applied Biosystems). Net
GUS activity of each strain was corrected for the basal fluorescence of P.
107 stewartii DC283 or E. coli SΦ200 carrying pL6GUSC without an insert.
-1 -1 -1 Specific activity was expressed in GUS units (pmol MU min OD600 ml of culture at 37°C). β-galactosidase activity was measured fluorometrically as described by Miller (1992) with the following modifications to adapt the protocol to the multiplate reader. Aliquots of cultures (100 µl) were mixed with 100 µl of a 2X β-galactosidase assay buffer (2.5 mM 4-methyl-umbelliferyl-β-D-galactopyranoside, 30%
DMSO, 200 mM NaCl in 200 mM potassium phosphate buffer, pH 7) and data collected as described above. β-galactosidase activity units were
-1 -1 -1 pmol MU min OD600nm ml of culture at 25°C.
2.2.11 Analysis of gfp reporter gene expression by flow cytometry
To quantify gfp expression, bacterial cells, collected by centrifugation
from growth media or from ooze extracted from plant tissues, were
resuspended and fixed in sterile filtered 0.5% paraformaldehyde at a final
OD600nm of 0.1-0.2. Suspensions were scanned in a FACS-Calibur
benchtop flow cytometer (Bencton-Dickinson) at the Ohio State
University Flow Core facility. The argon ion laser was tuned at 488 nm
and equipped with a 515/40 nm band-pass emission filter. GFP expression
was detected by forward and 90° side fluorescent light scattering. About
20,000 bacteria were counted per sample, Side scattering was
preferentially used to reduce the background noise. Data are reported as
distribution histograms of fluorescence vs. the number of cells (“events”). 108
2.3 RESULTS
2.3.1 Sequence analysis of the hrp regulatory region of P. stewartii
The hrp regulatory region was identified by transposon mutagenesis and
shown to contain three complementation groups (Frederick et al., 2001).
The region was subcloned from the cosmid pES1044 into two contiguous
plasmids, pRF8 and pRF201SK (Fig. 2.1). Prior sequencing of pRF8
(Frederick et al., 1993) revealed a single ORF, hrpS, which encodes an
NtrC-like enhancer protein homologous to HrpS from P. syringae pathovars (Rahme et al., 1991; Xiao et al., 1994), E. amylovora (Wei et al., 2000b) and P. agglomerans pv. gypsophilae (Mor et al., 2001). The main features of hrpS have been previously described (Frederick et al.,
1993). The sequence of the pRF201SK insert revealed three ORFs between hrpJ and hrpS, which have been designated hrpL, hrpX and hrpY. All of the ORFs in the regulatory region are transcribed from right to left as shown in Fig. 2.1. A detailed summary of the sequence analysis of the hrpL-hrpXY-hrpS region is shown in Table 2.3. Sequence analysis of the
54 region upstream of hrpL revealed a σ consensus box (CTGGCA-N6 -
TTGCT; probability score = 0.92) at -51 bp from the start codon of hrpL.
This was preceded by an A+T rich region containing putative integration
host factor (IHF) binding elements. A single dyad-symmetrical site
(TGCAA-N4 -TTGCA), weakly matching the NtrC-dependent upstream
activating site (TGCACC-N5 -GGTGCA; Porter et al., 1995), was located
109 at –163 bp. The predicted 21.3-kDa protein encoded by hrpL is
homologous to alternate sigma factors of the ECF σ 70 family (Lonetto et
al., 1994), including HrpL of E. amylovora (E value = 3e-74, 75%
identity; Wei and Beer 1995), P. agglomerans pv. gypsophilae (E = 2e-79,
80% identity; Mor et al., 2001) and P. syringae pathovars (E = 6e-30, 47
to 48% identity; Rahme et al., 1991; Xiao and Hutcheson 1994).
The hrpX ORF is located 144 bp downstream from the stop codon of
70 hrpL. A weak σ promoter consensus sequence (TGTGGA-N17-TAACAA,
probability score = 0.87) is located at -22 bp. Only 30 bp separates hrpX
from the downstream hrpY gene. The predicted proteins encoded by hrpX
(485 aa) and hrpY (213 aa) have homology to bacterial two-component
signal transduction systems, with HrpX similar to “orthodox” sensor
kinases and HrpY similar to bacterial response regulators (Hoch and
Silhavy, 1995). HrpX has two tandem PAS domains (Taylor and Zhulin,
1999) in the input region. HrpY belongs to the FixJ class of response
regulators with a conserved aspartate (D57) that is the putative site of
phosphorylation as inferred from multiple sequence alignments (not
shown). The closest PSI-BLAST matches in the GenBank database for these two genes were the ortholog HrpX/HrpY proteins of P. agglomerans pv. gypsophilae (E = 0, 79% identity for HrpX; E = e-103, 88% identity for HrpY; Mor et al.,, 2001) and E. amylovora (E = 0, 79% identity for
HrpX; E = 2e-96, 84% identity for HrpY; Wei et al., 2000b).
Interestingly, a putative HrpX and HrpY orthologous pair (E = 3e-40 for
110 HrpX; E = 9e-39 for HrpY) is also present in P. syringae DC3000
(Unfinished Microbial Genomes BLAST page at NCBI http://www.ncbi.nlm.nih.gov).
Our sequence analysis did not reveal any Rho-independent terminators
immediately following the end of hrpY, and multiple translational stops
are present in the 842-bp hrpY-hrpS intergenic region. BLAST searches of
the GenBank database indicated that a 483-bp IS-like element is present at
-23 bp from the start of hrpS (Fig. 2.2). This putative IS element is
flanked by two 8-bp direct repeats and contains two 75-bp imperfect
inverted repeats. The latter match sequences flanking unidentified
prophage genes and integrases present in the chromosomes and virulence
plasmids of Salmonella typhimurium and S. typhi (Genbank accessions
AE006471, AE008744, AL627268) and also flanking virulence ORFs 2
and 3 in P. syringae pv. phaseolicola plasmid pAV511 (accession
AF141883).Analysis of the secondary structure of the IS element using
the mFold algorithm revealed a strong hairpin with a free energy of
dissociation of –149.2 kcal mol-1 (Fig. 2.2). However, the IS-like element
in P. stewartii does not contain an ORF. Several putative σ 70 promoter
sequences are present upstream of and inside the IS-like element. PCR
analysis of 12 P. stewartii strains from our lab collection revealed that
this IS remnant was present in the same physical position in all strains
tested (Fig. 2.3) but not in a highly related strain of P. ananas.
111 2.3.2 Null mutant phenotypes and epistasis analysis
In a previous study, we isolated Tn5 and Tn3HoHoI insertion mutations in hrpL, hrpX, hrpY and hrpS and recombined them into the genome of P.
stewartii wild-type strain DC283, producing strains hrpL::Tn3HoHoI
(MEX105), hrpX::Tn3HoHoI (MEX116), hrpY::Tn5 (DM064) and
hrpS::Tn5 (DM758) (Fig. 2.1; Frederick et al., 2001). All of these mutants
have a Hrp- phenotype, cannot synthesize harpin and exhibit down-
regulation of several cosmid-borne hrp-lacZ fusions in secretion genes.
Since transposon mutations in hrpX are polar on hrpY, we constructed two
new nonpolar, in-frame, deletion mutants of hrpX, ∆hrpX(17-414) (DM2790)
and ∆hrpX(17-470) (DM2788) (Fig. 2.1). In tobacco, the newly constructed
∆hrpX mutants were unable to produce HR symptoms, even when
infiltrated at high cell densities (>109 CFU ml-1). On sweet corn
seedlings, mean disease severity ratings at 3 days after inoculation were
2.9 ± 0.2 and 1.9 ± 1.0 for DC283 and DM2790, respectively. Although
these means are not significantly different at P=0.05, we observed large
variations in both infection rates and symptom severity with the mutant
but not with the wild-type strain. In general, the ∆hrpX mutants were
capable of attaining wild-type levels of virulence, but with variable
response times and incidence. This may mean that the hrpX mutants are
much more sensitive to physiological differences between individual
seedlings.
112 Complementation analysis was done to determine the epistatic
interactions between different genes and rule out the possibility that some
phenotypes were due to second-site mutations. Low copy number plasmids
(pRK415 or pLAFR3) carrying hrpXY+, hrpY+, hrpS+ and hrpL+ (Fig. 2.1)
were constructed and introduced into different mutants. The ability of
these plasmids to restore HR elicitation in tobacco and virulence on sweet
corn to the mutants was tested (Table 2.4). In all cases where virulence
was restored, the complemented mutants were fully virulent (disease
severity rating ≥ 2.5 in at least two experiments) and the bacteria did not
need to be preinduced in IM to cause the HR. The Plac-hrpL plasmid,
pMM6, restored HR elicitation in tobacco and virulence on sweet corn to
all the insertion and deletion mutants in the regulatory region. Next, the
+ Plac-hrpS plasmid, pRF205, was able to restore the Hrp phenotype to
strains with mutations in hrpX and/or hrpY (strains MEX116, DM2788,
DM2790, DM064) and hrpS (DM758) and to a double hrpY hrpS mutant
(DM733), but not to an hrpL mutant (MEX105). Finally, plasmid pMM52, which contains Plac-hrpY, was able to restore HR elicitation and full virulence to ∆hrpX (DM2788, DM2790) and hrpY (DM064) mutants, but
not to hrpS (DM758) and hrpL (MEX105) mutants. Based on the
observations that i) mutations in hrpS were not suppressed by hrpY+, ii)
constitutive expression of hrpS+ or hrpL+ restored the Hrp+ phenotype to
hrpS and hrpY hrpS mutants, and iii) constitutive expression of hrpY+ and
113 hrpS+ alleles did not overcome the Hrp- phenotype of hrpL mutants, we
propose the following basic model for the Hrp regulatory cascade: hrpY Æ
hrpS Æ hrpL Æ hrp box promoters.
2.3.3 Expression of hrp regulatory genes in different genetic
backgrounds
To confirm the order of the regulatory cascade, hrpY::Tn5, hrpS::Tn5
and hrpL::aphA3 insertions were introduced into the chromosome of
selected hrp-lacZ or hrp-uidA strains to create double regulatory mutants.
In addition, we tested the effects of constitutively expressing various
plasmid-borne regulatory genes on chromosomally located lacZ and uidA
transcriptional fusions to hrpX, hrpS, hrpL and hrpJ promoters. Low copy number plasmids carrying Plac-hrpY, Plac-hrpS and Plac-hrpL were used to
over-express these genes and β-galactosidase or GUS activity of the
reporter fusions was measured following induction in IM liquid medium.
Results from these experiments are summarized in Fig. 2.4.
A second-site mutation in hrpY abolished expression of hrpS-lacZ and
hrpL-uidA (Fig. 2.4C) and a mutation hrpS decreased expression of hrpL- uidA (Fig. 2.4B), as expected from the epistatic analysis. Over-expressing hrpY turned up the hrpX-lacZ fusion ca. two-fold (Fig. 2.4A), showing
that hrpXY is a potentially autoregulated operon. Plac-hrpY stimulated the
hrpS-lacZ fusion ca. 8-fold in a hrpY+ background and ca. 20-fold in a
hrpY hrpS-lacZ double mutant (Fig. 2.4B). Plac-hrpY also up-regulated
114 hrpL ca. 7-fold in the hrpY+ chromosomal background and ca. 5-fold in
the hrpY hrpL-uidA double mutant, but it repressed hrpL expression in the
hrpS hrpL-uidA double mutant (Fig. 2.4C). The inability of HrpY to
increase hrpJ expression in strains containing a second-site hrpS or hrpL mutation (strains DM701 and DM786) further indicates that it does not directly activate hrpLp or hrp-box promoters, respectively (Fig. 2.4D). In
+ + a hrpS hrpXY background, the Plac-hrpS plasmid up-regulated both hrpL-
uidA and hrpJ-lacZ fusions 12- and three-fold, respectively (Figs. 4C and
4D). This effect was even more apparent in the hrpS hrpJ-lacZ and hrpY
hrpJ-lacZ double mutants, where expression increased 44- and 82-fold, respectively (Fig. 2.4D). However, Plac-hrpS did not affect hrpJ
expression in a hrpL hrpJ-lacZ double mutant (Fig. 2.4D), demonstrating
that it does not directly affect genes with hrp box promoters. Finally, the
Plac-hrpL plasmid up-regulated hrpJ-lacZ fusions in all genetic
backgrounds (hrpXY, hrpY, hrpS and hrpL; Fig. 2.4D), thereby confirming
that hrpL is the last gene in the hrp regulatory cascade. Significantly, the
expression of the hrpJ-lac fusions in this experiment mirrored the
virulence of the same strains in the above mutational and epistasis
analysis.
Since the lack of apparent terminators following the hrpL and hrpXY
operons means that transposon-induced mutations could have partial polar
effects on the level of expression of downstream genes, we repeated the
above experiments with plasmid-borne, reporter gene fusions in trans to
115 the tested regulatory mutations. We therefore constructed a series of
reporter fusions in pPL6GUS. These plasmids contained the regulatory
regions upstream of the hrpX, hrpS and hrpL ORFs fused to a uidA
reporter gene with its own ribosomal binding site (Fig. 2.1). Following induction in IM liquid medium, expression of each promoter was assayed in wild-type strain DC283 and in mutants lacking a hrp regulatory gene
(Table 2.5). Transcription from the hrpXp region, between –198 and +46
bp from the hrpX start, was high and constitutive in DC283. Expression of the hrpS-uidA fusion, containing the region between –928 and +384 bp,
was reduced up to 10-fold in hrpY and ∆hrpX mutants as compared to
DC283. The hrpL-uidA fusion was reduced ca. 250-fold in hrpS, ∆hrpX
and hrpY mutants.
Since Wei et al., (2000b) proposed that HrpS and HrpY both directly
activate hrpL in E. amylovora, we recreated each step of the P. stewartii
regulatory cascade in E. coli SΦ200, a genetic background that would lack other known hrp-specific regulatory molecules. We constructed E. coli
strains that harbored one of the above low copy hrp-uidA reporter
plasmids along with a high copy number plasmid that over-expressed a
regulatory gene (plasmids pMM46, pRF8, pMM22 carrying hrpY, hrpS and
hrpL, respectively). These strains were grown in LB broth, the regulatory
gene was induced with IPTG, and the cells were assayed for GUS activity.
(The following data are expressed as means ± S.D. They from
representative experiments with at least three replicates each and
116 statistical significance P was analyzed by ANOVA). The hrpXp-uidA
fusion in pMM25 was constitutively expressed in E. coli, but was further
stimulated as much as 6-fold by over-expression of hrpY (from 26 ± 6 to
127 ± 10 units; P <0.05). The hrpS-uidA fusion carried by pMM50 was up-
regulated 8-fold by HrpY (from 5 ± 2 to 42 ± 5 units; P<0.05) and the
hrpL-uidA fusion in pDM2785 was activated by HrpS (30-fold from 79 ±
20 to 2370 ± 817 units; P<0.001). In contrast, over-expression of hrpY did
not enhance hrpLp expression, but instead it consistently repressed its
basal level by ca. 6-fold (from 79 ± 20 to 12 ± 3 units; P<0.001). Finally,
the hrpJp-uidA fusion, which contains a promoter with an hrp-box, was
strongly activated by hrpL expressed from plasmid pMM22 (2000-fold
induction from 2 to 3970 units; P<0.001).
Overall, our data support a model where HrpY autoregulates hrpXY
expression, stimulates hrpS transcription and represses hrpL. These
results further confirm that HrpS activates hrpL and then HrpL alone
controls expression of hrp box promoters.
2.3.4 Role of the conserved aspartate in HrpY
Based on sequence homology and protein modeling (analysis not
shown), the conserved aspartate residue at position 57 is the most likely
phosphorylation site in HrpY. To test if D57 is required for activation of
HrpY, we mutated this codon to alanine and asparagine. Both
substitutions were expected to prevent phosphorylation, while minimizing
117 structural effects on the protein, as shown for similar response regulators
(Jin et al., 1990; Moore et al., 1993; Hoch and Silhavy 1995). The
missense mutant alleles were constructed in vitro by spliced overlap
extension PCR (SOE-PCR) and exchanged for the wild-type locus in the
chromosome of DC283 using the positive selection, suicide vector pLD55.
Both the hrpY[D57N] and the hrpY[D57A] chromosomal mutants were
completely unable to infect corn whorls or to elicit the HR when
infiltrated into tobacco leaves; they were complemented for both
phenotypes by the Plac-hrpY plasmid, pMM52. Unexpectedly, low copy
number plasmids carrying the same mutant alleles, pMM74 Plac-
hrpY[D57A] and pMM118 Plac-hrpY[D57N], were also able to complement
the null phenotype of the D57A, D57N and Tn5 hrpY chromosomal
mutants. As discussed below, this type of overexpression artifact is
common with two-component regulators and necessitates the use of single
copy mutant alleles for assessing phenotypes.
2.3.5 Effects of environmental conditions on the expression of hrp genes in vitro
The hrp/wts genes of P. stewartii are repressed in rich media (i.e. media with complex nitrogen sources) and induced in IM minimal inducing medium (Coplin et al., 1992, Frederick et al., 2001). To determine if the promoters of all the hrp regulatory genes would behave similarly to media repression, we compared the expression of the plasmid-borne hrpJp-uidA,
118 hrpLp-uidA, hrpSp-uidA and hrpXp-uidA fusions in wild-type strain
DC283 strain grown in LB broth and IM pH5.5. The results showed that all of the uidA gene fusions were dramatically repressed (up to ca. 3000- fold) in LB broth except for hrpXp-uidA, which was constitutively expressed (Fig. 2.5). The same promoter regions were cloned into the reporter plasmid pPROBEgfp[AAV] to form gfp fusions. This new set of
gene fusions allows the study of gene expression in vivo. Here we report
preliminary results that represent a proof of concept for the use of
unstable gfp fusions in analyzing virulence gene expression in plants. The
basal activity of the gfp fusions was measured in LB using flow-
cytometry. Gene expression data are presented as histograms showing the
distribution of the intensity of side (90º) light scattering (“90ºLS Log”)
upon excitation of the labels at 488 nm. Growth in corn increased the
proportion of cells fluorescing above the threshold from 17 to 36 fold
(Fig. 2.6). Upshifts of the median values was not as dramatic. It could be
that at +3 DPI only a small fraction of bacteria are still interacting with
intact cells, whereas the majority has turned down the expression of the
type III secretion system.
The regulatory effects of pH, phosphate, ammonia, and sodium chloride
were tested in vitro using chromosomal hrpX-lacZ, hrpS-lacZ and hrpJ-
lacZ fusions (Fig. 2.7), to avoid copy number artifacts while analyzing
the effect of individual environmental signals. IM was used as a basal
medium to test each variable. Expression of hrpS-lacZ and hrpJ-lacZ was
119 decreased by neutral to basic pH values (with maximal expression at pH
5.5) and by high potassium phosphate, ammonium chloride and sodium chloride levels. When we plotted the dose-response curves for each salt, expressed as its osmotic pressure (calculated by approximating activities to concentration values), the curves overlapped (data not shown), implying that part of the ability of IM to induce hrp genes could be due to its low osmolarity per se, rather than to nitrogen or phosphate starvation.
In contrast, expression of hrpX-lacZ was not affected by pH and was only slightly reduced by addition of the three salts (Fig. 2.7). In rich culture media, plasmid pRF205 Plac-hrpS relieved repression of promoters
downstream of hrpS in the cascade (data not shown). This was consistent
with the ability of pRF205 to enable P. stewartii to elicit an HR in
tobacco irrespective of the media used to grow the inoculum. We also
tested the effect of 2 mM nicotinic acid on wild-type DC283 cells
carrying pDM2531 hrpJ-uidA during growth in IM pH5.5. Expression of
this fusion in IM was reduced 5-fold by nicotinic acid (from 7357 ± 138
to 1439 ± 60 units ± S.E. in two experiments).
2.4 DISCUSSION
Bacterial virulence factors are usually tightly regulated and controlled
by environmental stimuli (Finlay and Falkow 1997). Mediating these
signals are complex, often hierarchical, regulatory cascades that integrate
multiple inputs and finely tune the infection process. The cascades
120 controlling type III and type IV secretion systems in several mammalian
and plant pathogens have been shown to include complete or partial two-
component signal transduction pairs. These constitute a common strategy
for linking the expression of energy-consuming secretion apparatuses to signals indicating the presence of a host. In this study, we present
evidence for a regulatory cascade that activates hrp/hrc type III
secretion/effector genes in P. stewartii in response to in vitro stimuli that
mimic the host environment. The cascade involves, from the top down
(Fig. 2.8), a two-component signal transduction pair (HrpX and HrpY), a
transcriptional activator (HrpS), and finally an alternative sigma factor
(HrpL). In this study, we first confirmed that HrpS and HrpL serve the
same functions in P. stewartii as they do in E. amylovora and P. syringae
(Xiao et al., 1992; Rahme et al., 1991; Wei et al., 1995, 2000;
Hendrickson et al., 2000ab; Hutcheson et al., 2001) and then we focused on the role of the HrpX/HrpY two-component system.
The occurrence of orthologs of HrpX/HrpY in E. amylovora (Wei et al.,
2000b), P. agglomerans pv. gypsophilae (Mor et al., 2001), P.
chrysanthemum (Ham, J.H., Rojas, C.M., Collmer, A., unpublished;
GenBank accession no. AF448202) and possibly in P. syringae pv. tomato
DC3000 (Unfinished Microbial Genomes database, available on line at the
NCBI or TIGR web sites) suggests that this two-component system may
represent a common mechanism for the environmental regulation of group
I hrp genes. However, only the functional and genetic analysis of E.
121 amylovora hrpXY has been published (Wei et al., 2000b) to date. The role
proposed by Wei et al., (2000b) for E. amylovora HrpY is different from
the one we present here for P. stewartii in that HrpY appears to serve as a modulator of HrpS activation of hrpL, instead of activating hrpS. This conclusion was based on their finding that a nonpolar hrpY::TnphoA mutation did not affect a chromosomal hrpS::Tn5-gusA1 reporter and on an earlier observation (Wei et al., 1995) that a hrpS mutation only partially regulates hrpL. A similar, independent study on hrp gene regulation in P. agglomerans pv. gypsophilae has been published
concurrently with ours (Nizan-Koren et. al. 2002). Their results with
HrpY are in complete agreement with our findings for P. stewartii. Other key data in Wei et al., (2000b), such as the inability of hrpY to complement a hrpS mutation, the downregulation of hrpL in a hrpY background and the autoregulation of the hrpXY operon by HrpY, are consistent with both our results and those in P. agglomerans pv.
gypsophilae. This suggests that common regulatory elements may interact
differently in the pantoeas and some other erwinias than they do in E.
amylovora. An extreme example of this is found in P. carotovorum,
which has both hrpS and hrpL. However, hrpL expression in this species
is constitutive and the hrp genes are post-transcriptionally regulated by
RsmA, which affects the stability of hrpL mRNA (Chatterjee et al., 2002).
In view of the apparent differences in the role of HrpY between P. stewartii and E. amylovora, we verified our hypothetical model using
122 multiple approaches. In particular, the regulatory effect of HrpY on hrpS
transcription was demonstrated repeatedly by measuring plasmid-borne
reporter gene fusions in hrp regulatory mutants (Table 2.5) by using
complementation tests with plasmid-borne hrpY+ in hrpS-lacZ or
hrpY::Tn5 hrpS-lacZ strains (Fig. 2.4B), and by reconstructing individual
steps of the cascade in E. coli. Moreover, over-expression of hrpY+
repressed the hrpL-uidA reporter in both a hrpS background (Fig. 2.4C)
and in E. coli genetic reconstruction experiments, further indicating that
HrpY cannot activate hrpL. It is possible that repression of PhrpL by unphosphorylated HrpY could provide a way of keeping the basal level of hrp gene expression low under noninducing conditions and/or provide a mechanism to turn it back down late in infection. In chapter 4 we present biochemical evidence for the function of HrpY as activator of hrpS and repressor of hrpL.
In the course of our complementation experiments, we found that a Plac-
hrpY plasmid activated hrpSp in the absence of hrpX and complemented
polar hrpXY mutants. These observations raised the question as to whether
or not HrpY always needs to be phosphorylated in order to activate hrp
genes. To address this question, we carried out site-directed mutagenesis
of the conserved aspartate (D57) in HrpY, which is the putative
phosphorylation site, and found that P. stewartii mutants carrying either
hrpY[D57N] or hrpY[D57A] chromosomal alleles in single copy were
completely Hrp-. This result indicated that D57 is required for HrpY to
123 function as a positive regulator and is genetic evidence that phosphorylation is required for HrpY activation. We believe that the avirulence of these mutants reflects the role of D57 in phosphorylation, rather than any nonspecific structural changes brought about by the amino acid substitutions. This role is supported by the solvent-exposed position of this residue (based on 3D protein modeling; not shown), its conserved position and the type of substitutions performed as shown for other well studied receiver domains (Jin et al., 1990; Moore et al., 1993).
Consequently, we attribute our finding that plasmid-borne wild-type HrpY and D57 mutant alleles were active in the absence of hrpX to an over- expression artefact that is commonly encountered in genetic studies of two-component systems (Hoch and Silhavy 1995). For example, in E. coli, overexpression of UhpA (a related FixJ-class response regulator) activated its target promoter uhpTp. This over-expression effect was observed with both wild-type UhpA and a D54A mutant protein (Shattuck-
Eidens and Kadner 1983; Kadner 1995). Two common explanations for this phenomenon are cross-talk with paralog kinases and the ability of the unphosphorylated response regulator to bind promoters at abnormally high concentrations (Hoch and Silhavy 1995).
In general, hrp genes are induced in minimal media that mimic conditions found in planta (Lindgren 1997), but these can vary considerably between species. For example, hrp genes in E. amylovora are stimulated by low pH and repressed by glucose, ammonium salts,
124 asparagine, histidine and nicotinic acid, but are unaffected by osmolarity
(Wei et al., 1992). P. syringae hrp genes are likewise induced at low pH in minimal media, but they are nonspecifically repressed by high salt concentrations (Rahme et al., 1992). The results of our study showed that expression of many hrp genes in P. stewartii was similarly repressed by rich media, activated in planta and regulated by pH and nitrogen sources, but, in contrast to E. amylovora, they were also affected by salt concentration (Fig. 2.7). hrpS-lacZ and hrpJ-lacZ chromosomal fusions were inhibited at neutral to basic pH and at high concentrations of potassium phosphate, sodium chloride and ammonium sulfate. In a previous study, we found that nitrates are also strongly repressing, but casamino acids increase expression rather than repress it (Coplin et al.,
1992).
For our studies on the role of the HrpX/HrpY two-component system in
sensing environmental signals, we constructed two P. stewartii mutants
with different in-frame deletions within the kinase domain of HrpX. In these strains, the expression of hrpS, hrpL and hrpJ was greatly reduced
(to levels comparable to a null hrpY mutant) when assayed in IM (Table
2.4) and the ∆hrpX mutants could not cause an HR on tobacco, which in
P. stewartii requires pre-induction in IM. This is the first clear indication
in any system that HrpX is needed to sense the chemical/physiological
signals provided in vitro by an apoplast-mimicking Hrp-inducing medium.
The hrpXY operon itself maintained a high basal level of expression in
125 both rich media (Fig. 2.5) and in IM medium at neutral pH with added
salts (Fig. 2.7), whereas the hrpS promoter was repressed under these
conditions. Thus, hrpS appears to be the first gene in the regulatory
cascade that is under strong environmental control and its transcriptional
regulation by phosphorylated HrpY and global regulators (M. Merighi and
D. Coplin, unpublished) may therefore be the key point in the cascade
where environmental signals are integrated.
The exact nature of the signals detected by HrpX is currently under
investigation in our laboratory. At this point, however, the sequence
analysis of HrpX offers a possible explanation as to how the hrp regulon
is able to respond to such a wide range of environmental conditions. The
absence of transmembrane regions indicates that HrpX is probably a
cytoplasmic protein and the presence of duplicated PAS domains in the N-
terminal portion suggests that its signal may be internal. PAS domains are
versatile cytosolic protein modules involved in monitoring light, redox potential, oxygen, small ligands and the energy level inside cells (Taylor and Zhulin 1999; Stephenson and Hoch 2001). Cluster analysis of a
diverse set of PAS domains from prokaryotic sensor proteins revealed that the PAS domains of HrpX are most closely related to those of Azotobacter vinelandii NifL (analysis not shown), which is a flavoprotein with FAD as its prosthetic group (reviewed in Taylor and Zhulin, 1999). Therefore by analogy to NifL, HrpX might sense redox potentials and energy changes within the cell. This hypothesis would explain how the Hrp regulon is
126 able to respond to most of the conditions known to induce hrp genes, i.e
acidity, phosphate and nitrogen starvation, all of which could affect
cellular redox potentials.
The importance of HrpX during infection appears similar in
Pantoea/Erwinia species in that it is not absolutely required for virulence.
In E. amylovora, a nonpolar hrpX::TnphoA mutant was greatly reduced in
virulence, response time and growth in planta and the expression of hrpL was reduced five-fold in inducing medium (Wei et al., 2000b). In contrast, a nonpolar hrpX::mini-Tn10-Km mutant of P. agglomerans pv. gypsophilae remained fully virulent and did not show any decrease in downstream hrp gene expression in planta, but the effect of this mutation was not determined in an inducing medium (Nizan-Koren et al., 2002).
Since the basis for the nonpolarity of TnphoA and mini-Tn10 transposons
in Erwinia spp. is not understood, it is possible that hrpY was not
expressed at normal levels in these experiments. The two P. stewartii
∆hrpX mutants used in this study were only slightly reduced in virulence,
infectivity and response time, but they exhibited much greater variability
in symptom severity and incidence on corn seedlings. This suggests that
the ability of the pathogen to sense the nutritional status of its host is
very important to insure consistent infection. On the other hand, given
that ∆hrpX mutants still retained significant virulence on sweet corn,
while nonphosphorylatable mutants of HrpY were totally nonpathogenic,
we hypothesize that there is an alternative HrpX-independent pathway
127 leading to HrpY activation in planta. This could reflect either the increased accumulation of small phosphodonor molecules (McCleary et al., 1993) under starvation conditions or the action of an alternative bacterial kinase specifically activated in corn (but not in tobacco or IM).
It will be interesting to learn if this alternate phosphorylation pathway responds to host cell contact or specific plant inducer molecules.
2.5 SUMMARY
A regulatory cascade activating hrp/hrc type III secretion and effector genes was delineated in Pantoea stewartii subsp. stewartii, a bacterial pathogen of corn. Four hrp regulatory genes were characterized: hrpX and hrpY encode the sensor-kinase and response regulator, respectively, of a two-component signal transduction system; hrpS encodes an NtrC-like transcriptional enhancer; and hrpL encodes an alternative sigma factor.
Epistasis analysis, expression studies using gene fusions, and genetic reconstruction of each step in Escherichia coli were used to delineate the following pathway: HrpY activates hrpS and also positively autoregulates the hrpXY operon. In turn, HrpS is required for full activation of the σ 54- dependent hrpL promoter. Finally, HrpL controls expression of all known hrp and wts genes. In vitro, hrpS and all downstream hrp genes were regulated by pH and salt concentration. Mutants with in-frame deletions in hrpX were still partially virulent on corn, but unable to sense the chemical and/or metabolic signals that induce hrp genes in vitro. Site-
128 directed mutagenesis of HrpY indicated that aspartate 57 is the probable phosphorylation site and that it is needed for activity. These findings suggest that both HrpX and an alternate mechanism are involved in the activation of HrpY in planta.
129 Strain, or plasmid Relevant phenotype(s) and/or genotype(s) Source or reference
Escherichia coli
r q + + BW20339 F' 128::Tn10-12(Kan ) lacI ∆lacZM15 pro(BA) /DE3(lac)X74 uidA(∆MluI)::pir recA1
∆phoA532 ∆(phnC∆DEFGHIJKLMNOP)33-30 Metcalf et al., ( 1996)
DH10B F- mcrA ∆(mrr-hsdRMS-mcrBC) φ80dlacZ∆M15 ∆lacX74 endA1 recA1 deoR
∆(ara,leu)7697 araD139 galU galK nupG rpsL λ- Gibco BRL
- - - r HB101 F thi-1 hsd20 (r Bm B) sup E44 recA13 ara-14 leuB6 proA2 lacY1 rpsL20 (Sm )
xyl-5 mtl-1 Boyer and
Roulland-Dussoix (1969)
HB101Rif spontaneous Rifr strain Lab collection
Sφ200Rif metB strA purB ∆(agg-uidA-man) Rifr Wei et al., (2000)b
Pantoea stewarti subsp. stewartiii
SS104 Wild type, Wts+, HR+ ICPPBa
DC283 SS104 Nalr Coplin et al., (1986)
DC102 P. stewartii corn isolate A. Vidaver Lab
DC110 P. stewartii SF8-2. isolated in Urbana, OH Hooker, 1967
DC119 P. stewartii corn isolate S. Walker, 1976
DC145 P. stewartii ATCC29227 Lab collection
DC146 P. stewartii ATCC 29228 Lab collection
DC147 P. stewartii ATCC 29229 Lab collection
DC162 P. stewartii corn isolate, Hudson Valley, N.Y. Lab collection, 1975
DM064 DC283 hrpY1296::Tn5 (Kanr) Frederick et al., (2001)
DM701 DC356 ∆hrpS hrpJ268::Tn3HoHoI (Ampr, Rifr) This study
DM711 DC283 hrpJ79::Tn3HoHoI (Ampr) Frederick et al., (2001)
DM729 DC283 hrpY1296::Tn5 hrpJ79::Tn3HoHo (Kanr, Ampr) This study
DM733 DC283 hrpY64::Tn5 hrpS1::Tn3HoHoI (Kanr, Ampr) This study
DM758 DC283 hrpS224::Tn5 (Kanr) Frederick et al., (2001)
DM786 DC283 hrpJ79::Tn3HoHoI hrpL2560::aphA-3 (Kanr, Ampr) This study
DM2788 DC283 ∆hrpX(17-470) This study
DM2790 DC283 ∆hrpX(17-414) This study
DM2831 DC283 hrpL-uidA (Tetr) This study
Table 2.1. (continued)
130 Table 2.1 (continued)
DM2837 DC283 hrpY hrpL-uidA (Kanr Tetr) This study
DM2844 DC283 hrpS hrpL-uidA (Kanr, Tetr) This study
MEX1 DC283 hrpS1::Tn3HoHoI (Ampr) Frederick et al., (2001)
MEX105 DC283 hrpL105::Tn3HoHo (Ampr) Frederick et al., (2001)
MEX116 DC283 hrpX116::Tn3HoHoI (Ampr) Frederick et al., (2001)
MM211 DC283 hrpY[D57A] This study
MM254 DC283 hrpY[D57N] This study
SW2 P. stewartii corn isolate, Wooster, OH Lab collection, 1974
SW20 P. stewartii corn isolate, Fayette Co., KY Lab collection, 1975
SW21 P. stewartii corn isolate, Grayson Co., KY Lab collection, 1975
SW4 P. ananas corn isolate, Portsmonth, OH Lab collection, 1975
Plasmids pBluescript
KS and SK (+) ColE1 αlacZ (Apr) Stratagene pLAFR3 IncP cos αlacZ (Tcr) Staskawicz et al., (1987)
pRK415 IncP αlacZ (Tcr) Keen et al., (1988)
pVK100 IncP cos (Tcr) Knauf and Nester (1982)
pLD55 R6K ori, αlacZ, tetAR (Apr) Metcalf et al., (1996) pUC18K promoter/terminator-less aphA-3 cassette Menard et al., (1993) pR751 IncP, (Tpr, Sp/Smr) Ruvkun and and Ausubel (1981)
pSU19 pACYC187 replicon (Cmr) Bartolome et al., 1991
pPL6GUSC pLAFR6 derivative carrying a promoterless uidA gene (Tcr) Knoop et al., (1991) pPROBE-KT pVS1 derivative (KmR) carrying a promoterless gfpmut3 gene Miller et al.,, 2000 pPROBEgfp[AAV] pBROBE derivative carrying a promoterless gfp[AAV] Miller et al.,, 2000
+ - r r r pRK2013::Tn7 ColE1 mob traRK2 ∆ repRK2 repE kan::Tn7 (Tp , Sm , Sp ) Ditta et al., (1980)
pDM1296 pES1044 hrpY::Tn5 Coplin et al., (1992)
pDM2560 pRF201 hrpL::aphA-3 This study
pDM2756 pDM2785 insert in pPROBEgfp[AAV] to create a hrpL-gfp[AAV] fusion This study
pDM2760 pDM2531 insert in pPROBEgfp[AAV] to create a hrpJ-gfp[AAV] fusion This study
pDM2724 pMM25 insert in pPROBEKT to create a hrpX-gfp fusion This study
pDM2842 pBKS (+) with PCR-made hrpX allele deleted from aa 17 to 414 This study
Table 2.1 (continued) 131 Table 2.1 (continued) pDM2846 pBKS (+) with PCR-made hrpX allele deleted from aa 17 to 470 This study pDM2848 pLD55 with the 0.98 kb SstI-XhoI fragment from pDM2842 This study pDM2851 pLD55 with the 1.1 kb SstI-XhoI fragment from pDM2846 This study pES1044 cosmid clone in pVK100 containing left half of hrp cluster Coplin et al., (1986) pES411 cosmid clone in pVK100 containing right half of hrp cluster Coplin et al., (1992)
+ pMM6 pRK415 with a 0.6 kb BamHI-EcoRI hrpL PCR fragment, transcribed from Plac This study
+ pMM22 pBSK(+) with a 0.6 kb BamHI-EcoRI hrpL PCR fragment, transcribed from Plac This study
pMM46 pBKS(+) with a 0.7 kb HindIII hrpY fragment made by PCR, transcribed from Plac This study pMM52 pRK415 with the 0.7 kb BamHI-EcoRI hrpY+ fragment from pMM46, transcribed
from Plac This study
pMM57 pBKS(+) with a 0.7 kb HindIII hrpY[D57A] fragment made by PCR-SOE This study
pMM58 pRF201SK with the 1.7 kb HindIII fragment from pRF8 and containing hrpL+,
hrpXY+, hrpS+ This study
pMM74 pRK415 with the 0.7 kb HindIII hrpY[D57A] fragment from pMM74,transcribed from PlacThis study
pMM92 pBKS(+) with a 0.7 kb HindIII hrpY[D57N] fragment made by PCR-SOE This study
pMM118 pRK415 with a 0.7 kb HindIII hrpY[D57N] fragment from pMM92, transcribed from Plac This study pMM165 pMM50 insert in pPROBEgfp[AAV] This study pMM245 pLD55 with the 0.7 kb SstI-XhoI fragment from pMM57 This study pMM246 pLD55 with the 0.7 kb SstI-XhoI fragment from pMM92 This study pMM330 pSU19 with the 0.7 kb SalI-SstI hrpY+ fragment from pMM46 transcribed
from Plac This study pRF201 pLAFR3 with the 3.6 kb BamHI-HindIII fragment containing hrpL+, hrpX+ and hrpY+
from pES1044 Coplin et al., (1992) pRF201SK pBSK(+) with the 3.6 kb BamHI-HindIII fragment from pRF201 This study pRF205 pVK100 with the 1.7 kb HindIII fragment containing hrpS from pES1044 Frederick et al., (1993) pRF8 pBSK(+) with the 1.7 kb HindIII fragment containing hrpS from pRF205 Frederick et al., (1993) pDM2531 pPL6GUSC with a 0.9 kb BamHI fragment to create a hrpJ-uidA fusion This study pDM2785 pPL6GUSC with a 389 bp BamHI PCR fragment to create a hrpL-uidA fusion This study pDM2791 pPL6GUSC with a 1.1 kb HindIII-BamHI fragment from pES411 to create
a wtsE-uidA fusion This study
Table 2.1 (continued)
132
Table 2.1 (continued) pMM25 pPL6GUSC with a 244 bp HindIII-BamHI PCR fragment to create a
hrpX-uidA fusion This study pMM50 pPL6GUSC with a 1.3 kb BamHI PCR fragment to create a hrpS-uidA fusion This study
Table 2.1. Bacterial strains and plasmids.
133
Primer Sewuence (5’Æ3’) Coordinates Source pBluescript and pPL6GUSC vectorsb
LacF ACCATGATTACGCCAAGCTC 813-794 This study
LacR GGCGATTAAGTTGGGTAACG 577-558 This study
5GUS CATTTCACGGGTTGGGGTTTCT 15-36 Jefferson (1987)
P. stewartii subsp. stewartiib
XFH ACGGAAGCTTACTTCAAGGAGGCGAGATGC 1364-1383 This study
4XRH ACGGAAGCTTGTCCTTCGCGCATGATGACG 2924-2905 This study
YFH CCGAAGCTTTTCGATAACAATATGGATAACAC 2855-2877 This study
YRH CCGAAGCTTATTAAGAGCAATCCTAAGAGA 3521-3501 This study
LFB CGCGGATCCACATTAAGCCAAACGGCAAAA 662-642 This study
LRE CCGGAATTCTGTTTTACCCCGTTCAGTT 1255-1237 This study
201.3 CCTGGCGAACCTCATACAGG 411-430 This study
201.1 CGCTATCATCTCGCCCTGTG 1691-1672 This study
YD57A TCTTTTACTGCTAGCTATGTGTATGCCTGG 3022-3051 This study
YD57N CAGGTCGATCTTTTACTTTTAAATATGTGTATGCCTGG 3014-3051 This study
YR2 GTTTCAGGATCGTTATCTTTCGTGATG 3204-3178 This study
EdX1 CGCGAATTCTTCGGTATTGCCCTGAACCT 919-938 This study
BdX1 CGCGGATCCTTTGATTGGCGTATCACTGCT 1427-1407 This study
BdX2 CGCGGATCCAACGAGGTTCTCATAGAGGT 2622-2641 This study
BdX3 CGCGGATCCACCAATGGAACCCGTGTGACT 2794-2813 This study
XdX3 CGCTCTAGATTTGCTCGGCGAGCATTTGG 3272-3253 This study
XFHG CCCAAGCTTACGTTCGCGTTTGTCACGGA 1182-1201 This study
XRBG CGCGGATCCTGATTGGCGTATCACTGCTA 1425-1406 This study
SFBG CGCGGATCCATAAAGCCCGCCTGATGGAA 3423-3442 This study
SR3BG CGCGGATCCTCTAGATGGCATGCTATCGATCTCGT 4735-4715 This study
JFBG CGCGGATCCGAGCTCTTTCCCAGGCAACAGAGTTAA 481-501 This study
LRBG CGCGGATCCTCTAGAATGACTTCCAGCCAGGTCAT 869-850 This study
ISR CGCGGATCCTCGGAATGAGTGCTCATTAT 4370-4389 This study
Table 2.2. (Continued)
134
Table 2.2 (continued)
a Positions of LacF/R and 5GUS primers are according to GenBank accession X52331
(pBluescriptKS+) and U12639 (uidA locus) respectively; positions for P. stewartii
primers correspond to GenBank accession AF282857 and do not include the 5’-end
restriction sites and anchoring tail. b Underlined bases represent engineered restriction sites; mutant codons are shown in
bold
Table 2.2. Primers
135
Regulatory elements/Protein features Protein homologies (E value; % similarity; % identity)b Accession # hrpL gene RBS (ACGGAG) at -6 54 σ promoter (CTGGCA-N6-TTGCT) at –51 IHF binding site (A+T rich region) at –55 UAS (TGCAA-N4-TTGCA) at –163 No terminators at 3’-end 21 kDa ECF sigma factor P.agglomerans pv. gypsophilae HrpL (2e-79; 80%; 89%) AAF76211 E. amylovora HrpL (1e-74; 75%; 86%) AAA91801 P. chrysanthemi HrpL (8e-49; 54%; 72%) AAM46690 P. syringae pv. tomato HrpL (6e-33; 48%; 66%) BAA82036 E. coli σE (3e-11; 28%; 50%) P34086 hrpX hrpY operon RBS (AGGAG) at -5 70 σ promoter (TGTGGA-N17-TAACAA) at –22 No terminators at 3’-end
HrpX, 56 kDa sensor kinase E. amylovora HrpX (0; 79%; 89%) AAD24682 No transmembrane domains P.agglomerans pv. gypsophilae HrpX (0; 72%; 85%) AAF76212 PAS-1 input domain (aa 16 to 65 P. chrysanthemi HrpX (e-146; 60%; 74%) AAM46691 PAS-2 input domain (aa 143 to 198) P. syringae pv. tomato DC3000 TCSc (3e-34; 31%; 50%) TIGR_317_5668 His-kinase domain (aa 296 to 485) X. campestris pv. campestris (1e-30; 30%; 49%) AAM41248 H296 putative phosphorylation site
HrpY: 24 kDa response regulator P.agglomerans subsp. gypsophilae HrpY (e-103; 88%; 93%) AAF76213 Receiver domain (aa 1 to 118) E. amylovora HrpY (6e-96; 83%; 89%) AAD24683 D57 putative phosphorylation site Pectobacterium chrysanthemi HrpY (8e-77; 70%; 80%) AAM46692 Output domain (aa 149 to 213) P. syringae pv. tomato DC3000 TCRc (3e-51; 49% ; 69%) TIGR_317_5668 HTH domain (LuxR homolog; aa Pectobacterium carotovorum ExpA (3e-39; 32%; 56%) CAA64809 69 to 194) Salmonella typhimurium SirA (2e-98; 33%; 58%) AAC44801 E. coli UhpA (2e-26; 33%; 55%) BVECAU
hrpS gene RBD (AGGGGC) 483-bp IS like at –23 Rho-independent terminator at 3’ end
36 kDa NtrC-like response regulator No N-terminal receiver domain E. amylovora HrpS (e-146; 79%; 90%) AAD24684 σ54-dependent ATPase domain P.agglomerans pv. gypsophilae HrpS (e-145; 79%; 91% AAF76214 C-terminal HTH domain Pectobacterium chrysanthemi HrpS (e-101; 59%; 73%) AAM46693 P. syringae pv. phaseolicola HrpS (3e-83; 57%; 70%) CAA54728 P. syringae pv. phaseolicola HrpR (2e-81; 56%; 70%) CAA54727
Table 2.3. (continued)
136 Table 2.3. (continued)
a Putative DNA regulatory elements, homologous proteins and conserved protein
domains were identified by computerized sequence analysis as described in Material
and Methods. RBS = ribosomal binding site; IHF = integration host factor; UAS=
upstream activating sequence (weakly matches NtrC UAS described by Porter et al.,
1995); HTH= helix-turn-helix motif; IS= insertion sequence; TCS = two-component
sensor; TCR = two-component regulator b Within parenthesis are the following Genbank statistics (in order): E value based on the July 8 2002 version of the Genbank nr database; % identity and % similarity (= positives) for BLASTP local alignments. c P. syringae pv. tomato. The hrpX hrpY homologs found by TBLASTP search form a
two-gene operon in the non-annotated contig TIGR_317_5668, recently renamed
PSPTO0559 and PSPTO0558 (Buell et al., 2003). The HrpX homolog in P. syringae
has the same domain organization of P. stewartii HrpX except for a periplasmic
binding protein domain in the N-terminal region (sequences courtesy of The Institute
for Genomic Research).
Table 2.3. Analysis of DNA and protein sequences
137
,
-hrpY lac -hrpL lac , pMM52 P - - - - hrpY + Path HR Path P hrpX hrpS
lac- HR n in IM pH 5.5 overnight.
Path P
hrpY no symptoms; +++ = many lesions; V variable a
lac- HR regulatory gene regulatory , - = no HR. Cells were grow
trans P + Path
HR hrpX uppression tests were low-copy number replicons: pDM1296 number replicons: were low-copy tests uppression
regulatory mutants and complementation analyse ys after inoculation of the whorls: - = hrp -hrpL + +++ + +++ + +++ + +++ + +++ + +++ + +++ + lac c V Path b ------+ +++ + +++ + +++ - + ------+ +++ + +++ + +++ + +++ + +++ + - - - -
------, and pMM6 P -hrpS lac HR + +++ + +++ + +++ + +++ + +++ + +++ + +++ + +++ + +++ + None 5 ------+ +++ - + - - - - - 5 3 - +++ 3 Tn ::Tn ::Tn ::Tn hrpX ∆ wild-type hrpS hrpY:: hrpJ hrpL c. Disease ratings on corn leaves at 3 da pRF205 P pRF205 a. complementation/s used for the plasmids All b. Hypersensitive response in tobacco. HR ratings: + = full ______DM758 type and of wild 2.4. Pathogenicity Table DM064 DC283 DC283 DM2790 _ MEX1 DM711
Strain Genotype
138
GUS activity (Units ± S.E.)c in different genetic backgroundsd Plasmid / Promoter fusionb Wild type ∆hrpX hrpY hrpS hrpL
pMM25 -198/+46 hrpXp 307±53 370±10 206±2 ND ND pMM50 -928/+384 hrpSp 135±9 20±9 21±4 119±12 ND pDM2785 -207/+182 hrpLp 6053±959 164±5 67±12 36±1 ND
pDM2531 -516/+425 hrpJp 4851±704 94±13 17±5 3±1 0±0 pDM2791 –914/+22 wtsEp 399±125 7±5 1±1 0±0 4±2
a Strains were grown in IM pH 5.5 broth to late exponential-early stationary phase and assayed for GUS activity as described in Material and Methods. b Transcriptional fusion names are preceded by the promoter fragment coordinates in bp (+1 is first base of ORF). c -1 -1 -1 One unit = 1 pmol MU min OD600 ml of culture at 37°C. S.E.= standard error of
two to six independent experiments, each with at least two replicates. d Strains: wild type = DC283; ∆hrpX = DM2790 (mutant DM2788 gave similar
results); hrpY = DM064; hrpS = DM758 or MEX1; hrpL = MEX105.
Table 2.5. Effects of hrp regulatory genes on plasmid-borne hrp::uidA
transcriptional fusions expressed in different genetic backgrounds.
139
Figure 2.1. Physical map of the regulatory region of the P. stewartii DC283 hrp gene cluster showing the hrpL, hrpX, hrpY and hrpS subclones used in this study.
Open boxes represent regulatory ORFs, the cross-hatched box is hrpJ, a secretion
operon, and the solid box represents an IS remnant in the hrpS 5’ regulatory region.
The direction of transcription is shown by arrows. Solid circles indicate Tn5
insertions, the open circles are point mutations, flags indicate Tn3HoHoI insertions
and their orientation, squares denote aphA-3 insertions, and ∆ indicates in-frame
deletions. Open triangles in front of genes represent putative σ 70 promoters, the
closed semicircle is a σ 54 promoter, and the open semicircle is a hrp box promoter.
P lac indicates the position of the constitutive lac promoter in the plasmid vector. Solid boxes labeled “uidA” indicate transcriptional fusions to the GUS gene in pL6GUSC.
Restriction enzyme abbreviations: H, HindIII, B, BamHI.
140
141
Figure 2.2. Analysis of the secondary structure of the IS element remnant located in front of hrpS.
The secondary structure corresponding to the transcript spanning the 483-bp IS-
element located in front of hrpS was performed using the Genebee server
(www.genebee.msu.su). An energy minimize 2D-folding model is shown. The
calculated free energy of folding was –149.2 Kcal mol-1.
142
1 2 3 4 5 6 7 8 9 10 11 12 13 14
1 kb
Figure 2.3. PCR detection in a collection of P. stewartii strains of the abortive IS element preceding hrpS.
Primers ISR and SFBG flanking the 483-bp IS-like element in front of hrpS were used to amplify by “colony PCR” a ca. 1-kb product from a collection of Pantoea stewartii strains. Only strain SW4 tested negative and was later identified as Pantoea ananas by biochemical and nutritional assays. Lane 1, 1-kb (+) molecular weight ladder; lane 2, P. stewartii strain SW2; lane 3, P. stewartii strain SW20; lane 4, P. stewartii strain DC102; lane 5, P. stewartii strain DC110; lane 6, P. stewartii strain
DC119; lane 7, P. stewartii strain DC145; lane 8, P. stewartii strain DC146; lane 9,
P. stewartii strain DC147; lane 10, P. stewartii strain DC162; lane 11, P. ananas strain SW4; lane 12, P. stewartii strain SW21; and lane 13, P. stewartii strain DC283.
143
Figure 2.4. β-galactosidase (β-Gal) and β-glucuronidase (GUS) activity of chromosomal hrp-lacZ or hrpL-uidA fusions in different P. stewartii genetic backgrounds, with and without various plasmid-borne Plac-hrp regulatory genes.
Cells were grown in IM pH 5.5 broth to late log phase and β-Gal specific activity
-1 -1 -1 was measured (1 unit = 1 pmol MU min OD600 ml of culture at 25°C). The first
column indicates which regulatory gene was constitutively expressed in trans (if any);
the second column indicates the second hrp mutation (if any) in the strain. Regulatory
gene clones: pMM52 Plac-hrpY and pMM46 Plac-hrpY (panel C; low copy pMM330 was
used as well with similar results); pRF205 Plac-hrpS (panel D) and pRF8 Plac-hrpS
(panel C); and pMM6 Plac-hrpL (panel D). Controls consisting of the vector plasmids
without inserts had no effect on expression of any of the gene fusions.
Panel A: β-Gal activity in hrpX-lacZ strain MEX116; Panel B: β-Gal activity in the
hrpS-lacZ strain MEX1 and hrpY hrpS-lacZ strain DM733. Panel C: GUS activity in
hrpL-uidA strain DM2831, hrpY hrpL-uidA strain DM2837 and hrpS hrpL-uidA strain
DM2844. Panel D: β-Gal activity in the hrpJ-lacZ strain DM711, hrpY hrpJ-lacZ
strain DM729, hrpL hrpJ-lacZ strain DM786, and ∆hrpS hrpJ-lacZ strain DM701.
144
145
10000
1000 rich medium 100 IMpH5.5 10 specific GUS activity (pmol MU min-1 OD-1) 1 DC283/pMM25 DC283/pMM50
DC283/pDM2531 DC283/pDM2785
Figure 2.5. Expression of plasmid-borne hrp-uidA fusions in P. stewartii DC283.
P. stewartii strain DC283 carrying hrp-uidA plasmid-borne fusions was grown in LB broth and IM pH5.5 liquid medium and GUS activity was measured using a fluorometric protocol as described in Materials and Methods. One unit = 1 pmol MU
-1 -1 -1 min OD600 ml of culture at 37°C. Error bars represent standard deviations from single experiments with at least two replicates. Strains: wild type = DC283; plasmid pMM25 contains -198/+46 hrpXp, pMM50 contains -928/+384 hrpSp, pDM2785
contains 207/+182 hrpLp, and pDM2531 contains -516/+425 hrpJp.
146
Figure 2.6. Flow-cytometry analysis of hrp-gfp fusion expression in P. stewartii
DC283 cells from infected corn plants compared to their expression in LB.
P. stewartii DC283 cells carrying hrp-gfp plasmid-borne fusions were either grown in
LB broth or recovered as ooze from the surface of inoculated in corn seedlings. The whorls of 8-day-old potted sweet corn seedlings were inoculated with bacteria supsended in 0.2% Tween-40. After two days the tops of the plants were cut off, placed in moist chambers and incubated for an additional 24 h; bacterial ooze was then collected from the leaves. Bacteria were washed in sterile saline solution and resuspended in 0.5% paraformaldehyde to fix the cells. Samples were analyzed in a benchtop flow cytometer fitted with an argon laser tuned at 488 nm. Side Light scattering fluorescence (“90ºLS Log”) of ca. 20,000 cells per sample was read at
515/40 nm band-pass filter. Gene expression values are reported as % of gated cells above the fluorescence threshold (M) in LB vs. corn. Red solid histogram = cells grown in LB. Black histogram: cells collected corn leaves at 3 DPI; pDM2724 is hrpX-gfp; pMM165 is hrpS-gfp[AAV]; pDM2756 is hrpS-gfp[AAV]; pDM2760 is hrpJ-gfp[AAV]
147
M=0.73% vs. 20% M=0.7% vs. 17.9%
M M
pDM2724 pMM165
M=0.7% vs. 12% M=0.7% vs. 24.9%
MM
pDM2756 pDM2760
M=% of cells above the fluorescence threshold upon growth in LB vs. in planta
148
Figure 2.7. Effect of environmental conditions on the expression of various hrp-
lacZ fusions.
The effects of neutral pH, 50 mM sodium chloride, 25 mM ammonium sulfate and
25 mM potassium phosphate are shown as a percentage of the activity in standard IM
pH 5.5 liquid medium (control). Cells were grown in modified IM, with one
component changed as indicated in the Z-axis of the graph, for 16 h and β-Gal activity
was measured. hrpX-lacZ = strain MEX116, hrpS-lacZ = strain MEX1 and hrpJ-lacZ =
DM711.
149
Figure 2.8. Working model for the hrp regulatory cascade in P. stewartii.
The HrpX sensor perceives undefined metabolic and/or stress signals and activates
HrpY via phosphorylation (indicated by ~P). A parallel phosphorylation mechanism
may also activate HrpY in planta. HrpY then activates hrpS and its own operon; HrpS
and σ 54 activate hrpL; and finally HrpL enables RNA polymerase core enzyme (E) to activate promoters containing hrp boxes (e.g. the harpin, wts and hrp secretion genes).
150
CHAPTER 3
A NOVEL AUTOREGULATORY LOOP ACTIVATED BY HRPS
FROM THE HRPL PROMOTER MODULATES EXPRESSION OF THE
P. STEWARTII TYPE III SECRETION SYSTEM REGULATORS
HRPXY, HRPS AND HRPL
3.1 INTRODUCTION
Pantoea stewartii subsp. stewartii is a bacterial pathogen of sweet corn and maize that causes wilting and leaf necrosis, preceded by a water- soaking phase (Pepper, 1967). Although extracellular polysaccharide
(EPS) is a major virulence factor contributing to wilting symptoms (Dolph et al., 1988; Coplin and Majerczak, 1990), the Hrp type III secretion system is absolutely required for growth in the host to high cell density
(>108 CFU/g tissue) and for development of both lesions and wilt symptoms (Coplin et al., 1986; Coplin et al., 1987). hrp genes are organized in horizontally mobile, pathogenicity islands (PAI) (Hutcheson,
1999; Kaper and Hacker, 1999) located, in different pathogens, either on the chromosome (Erwinia amylovora, Pectobacterium carotovorum,
Pectobacterium chrysanthemi, Pseudomonas syringae, Xanthomonas 151 campestris, Xanthomonas vesicatoria) or plasmids (Pantoea agglomerans
pv. gypsophiliea, Ralstonia solanacearum) (Rosenberg et al., 1982;
Boucher et al., 1987; Valinsky et al., 1998; Ezra et al., 2000). Hrp PAIs usually carry their specific regulatory genes linked to the secretion/effector loci, but additional unlinked genes are often required for hrp/hrc expression. All pathogens carrying group I hrp clusters (P.
syringae and erwinias) require the chromosomally encoded, nitrogen
starvation sigma factor σ 54 (RpoN) for expression of hrp secretion and
effector genes (Frederick et al., 1993; Hendrickson et al., 2000).
Furthermore, expression of P. carotovorum hrpL is reduced in a himA
(IHF) E. coli mutant (Chatterjee et al., 2002), implying a positive role for
IHF in hrp expression. On the other hand, in group II pathogens, X. vesicatoria hrpX, which encodes an AraC-like activator, is unlinked to the main PAI (Wengelnik and Bonas, 1996) and the OmpR-class response regulator HrpG lacks a physically linked cognate sensor kinase
(Wengelnik et al., 1996).
During the homing process for horizontally acquired DNA (Kaper and
Hacker, 1999), it is indeed reasonable to envisage the fortuitous interaction between regulatory elements in the PAI and various housekeeping regulatory genes already present in the recipient strain.
Some of these new interactions may mimic those present in the donor
152 strain, whereas others may be novel and selected for or against. This may adapt the expression pattern of the virulence genes to fit the new host environment.
Regulation of group I hrp clusters always involves three common regulatory proteins, HrpS, HrpL and RpoN, and probably IHF as well. As mentioned above, IHF regulates hrp genes in P. carotovorum, but its role is only assumed for the other group I pathogens, based on the presence of similar A+T rich regions 60-80 bp upstream of hrpL (Hutcheson, 1999;
Merighi et al., 2003). The Lon protease may act as an additional negative regulator of hrp gene expression as shown by a recent study on
Pseudomonas syringae (Bretz et al., 2002) and work from our lab (Ham and Coplin, unpublished) on P. stewartii. Phytobacterial pathogens of the erwinia group also use the HrpX/HrpY two-component system, which is not found in P. syringae.
In spite of the high similarity (around 80% identity) of the regulatory proteins, the 5’- and 3’-end region of their genes are quite different. This divergence is reflected in different basal levels of expression and different epistatic interactions of the regulatory genes in the various species. For instance, in E. amylovora, Wei and colleagues concluded that
HrpS is environmentally regulated in a HrpX/HrpY-independent manner and that both HrpY and HrpS act additively to control hrpL (Wei et al.,
2000b). On the other hand, the role of HrpX/HrpY in Pectobacterium spp.
153 is unclear, because hrpS expression appears to be constitutive (Chatterjee et al., 2002) and the phenotype and regulatory effects of polar hrpXY
mutations are not known.
In P. stewartii and P. agglomerans pv. gypsophiliae, HrpY has been
shown in genetic reconstruction experiments to activate hrpS, but not
hrpL. Moreover, in a E. coli rpoN+ himA+ background, HrpS alone appears
to be sufficient for full activation of hrpL (Chapter 2; Merighi et al.,
2003; Nizan-Koren et al., 2003). A significant difference between P.
stewartii and the other erwinias is the abortive IS element located just
upstream of the RBS of hrpS in P. stewartii (Merighi et al., 2003). This element might form a strong hairpin-like secondary structure that could
potentially alter expression of hrpS itself. In addition, it could also carry
additional regulatory elements that may explain some of the unique features of hrpS regulation in P. stewartii, e.g. its regulation by quorum
sensing as discussed in Chapter 6.
In all erwinias, the hrpL, hrpXY and hrpS regulatory operons are
colinear and transcribed in the same direction, from hrpL to hrpS (Fig. 1),
whereas in P. syringae, hrpL and hrpRS are located on opposite ends of
the hrp/hrc cluster, probably due to a translocation event during homing of the PAI (Hutcheson, 1999). In P. stewartii, differently from other erwinias, preliminary results indicated that hrpS was autoregulated in some manner. Ectopic expression of hrpS upregulated hrpS::lacZ and
hrpX::lacZ fusions, but the effect on hrpS was abolished by a polar 154 insertion in hrpY (Coplin, unpublished observation). The particular
transcriptional organization of the three regulatory operons and the
apparent absence of potential terminators downstream of hrpL and hrpY
(Merighi et al., 2003) suggested that transcription from either or both
PhrpL and PhrpX could continue into hrpS. Alternatively, increased
expression of hrpXY and trans-activation of hrpS by HrpY could also
cause this autoregulatory effect. Both scenarios would create a novel
autoregulatory loop in the regulatory pathway.
The purpose of this study was to analyze the molecular basis of the
positive autoregulatory loop triggered by HrpS and to identify the
regulatory elements, promoters and transcripts involved. In particular we
wanted to determine: (i) if the HrpS-dependent autoregulatory effect is
triggered by activation of the hrpL and/or hrpXY promoters; (ii) whether
autoregulation acts by the effect in trans of HrpY on hrpS or it is instead
due to the cis-effect of transcription continuing from hrpXY and/or hrpL
into hrpS.
We initially studied the effect of ectopic expression of hrpS in double
mutants with a chromosomal polar insertion in one regulatory gene and a
reporter gene fusion in another. After we defined the candidate promoters
involved in hrpS autoregulation, we characterized them by analyzing the
5’-ends of the various mRNAs and then we utilized this information to
demonstrate inter-operon transcription using RT-PCR. The existence of
155 long transcripts starting from hrpL and spanning hrpL and hrpXY and part
of 5’ hrpS was further shown by Northern blot analysis. The possible
roles of hrpS autoregulation in P. stewartii biology are discussed.
3.2 MATERIALS AND METHODS
3.2.1. Bacterial growth and media
The bacterial strains and plasmids used in this study are given in Table
3.1. Strains were grown in Luria-Bertani (LB) broth and agar (Ausubel et
al., 1987) for inoculum preparation and cloning procedures or in IM
liquid medium at pH5.5 (IM5.5) (Frederick et al., 2001) for induction of
hrp gene expression as described (Merighi et al., 2003). Selection for tetracycline sensitive strains was done in the fusaric acid medium
(Buchner selection) described by Metcalf et al. (1996). When required, antibiotics were added at the following concentrations: tetracycline at 20
µg/ml, nalidixic acid at 10 µg/ml, kanamycin at 50 µg/ml,
chloramphenicol at 50 µg/ml, and ampicillin at 200 µg/ml.
3.2.2 Molecular genetics and cloning techniques
Plasmids were transferred among strains by biparental mating or
triparental mating using pRK2013::Tn7 as helper plasmid (Ditta et al.,
1980) or introduced by electroporation. Double mutants were constructed
by allele exchange of mutations cloned in the suicide vector pLD55 with
positive selection of double recombinants in fusaric acid media as 156 described previously (Metcalf et al., 1996; Merighi et al., 2003).
Preparation of cell lysates, DNA purification, molecular cloning
experiments, Southern blot analysis and PCR reactions were performed
following standard procedures (Ausubel et al., 1987).
3.2.3 Construction of polar hrpL interposon mutants and of plasmid
pMM191
To construct a polar mutation in hrpL to use for allele exchange in P.
stewartii, the interposon Ω/Spr was purified as a 2 kb SmaI fragment from
plasmid pHP45::Ω and ligated in the NvuI site of plasmid pDM2533,
which contains the hrpL gene in a 1.3 kb PCR fragment. The resulting
plasmid, pDM2862, was digested with KpnI/SpeI and the 3.3 kb hrpL::Ω
fragment was subcloned into the suicide plasmid pLD55 to create
pDM2868. E. coli strain BW20339 (pDM2868) was crossed with strains
MEX1 hrpS-lacZ and MEX116 hrpX-lacZ by triparental mating (Frederick et al., 2001) and transconjugants were selected for Tetr. Single cross-over
mutants were streaked on fusaric acid medium to select for the second
recombination step as previously described (Metcalf et al., 1996).
Putative Tets double-recombinants strains were screened by PCR and
named DM2824 hrpL::Ω hrpS-lacZ and DM2830 hrpL::Ω hrpX-lacZ.
Interposon-chromosome junctions were confirmed by Southern blot
analysis. Plasmid pMM191, containing a 5’deletion of the hrpL regulatory
157 region, was constructed by amplifying a 315 bp fragment of hrpL with
primers LRBG and LFU and pMM58 as template DNA (Table 3.2). The
insert was cloned as a BamHI insert in pPL6GUSC and sequenced.
3.2.4 Analysis of β-glucuronidase or β-galactosidase activity
β-glucuronidase and β-Galactosidase activities in bacterial cultures
were measured using the fluorogenic substrates 4-methyl-umbelliferyl (4-
MU)-glucuronide (MUG) and 4-MU-galactoside (MUGal), respectively.
Assay were done in 96-well microtiter plates following standard protocols
(Miller, 1992; Jefferson, 1985) modified as previously described (Merighi
et al., 2003). One unit of enzyme activity is equal to 1 pmol 4-methyl-
umbelliferone (MU) per min per OD600.
3.2.5 Total RNA isolation
Total bacterial RNA was isolated from 5 ml of overnight cultures of
cells grown in IM pH5.5 liquid medium or 5 ml of exponentially growing
cultures in LB broth. Cells were pelleted and resuspended in 0.1 ml of
TEN100 buffer (10 mM Tris-HCl, 100 mM NaCl, 1 mM EDTA, pH 7.0).
Five volumes of Trizol (Invitrogen) at 65°C were added to the
resuspended cells, which were then vortexed, shaken at 65°C for 10 min
and centrifuged at 10,000 x g for 30 min at 4°C. The upper aqueous phase was extracted with 0.1 ml of water-saturated chloroform and the RNA was precipitated by adding one volume of isopropanol and centrifuging as 158 above. RNA was resuspended in 50 µl of DEPC-treated water and treated with DNase I (Invitrogen) according to the manufacturer’s instructions.
To assess the quality of the RNA preparations, 10 µl aliquots were analyzed by standard agarose gel electrophoresis with ethidium bromide staining.
3.2.6 Primer extension and DNA sequencing
Analysis of 5’-ends of transcripts was performed by primer extension
(Sambrook et al., 1989). Primers of the “PE” series described in Table 3.2 were labeled with ([γ32P] ATP and cDNA synthesis was carried out using
M-MLV reverse transcriptase (Promega) with 50 µg of RNA per reaction following the manufacturer’s instructions. DNA sequencing was performed with the same primers used for cDNA synthesis following the fmolTM cycle sequencing protocol from Promega. The labeled extension products were visualized by autoradiography with Kodak XOMAT films after electrophoresis in 6% denaturing acrylamide gels at 55 Watts in a
Bio-Rad sequencing apparatus. For weakly labelled cDNAs, a second exposure was performed, and the DNA ladder and cDNAs were merged electronically using Photoshop Elements (Adobe).
3.2.7 RT-PCR analysis
Analysis of inter-operon transcripts was performed using the Titan-One-
Tube RT-PCR protocol from Roche. Sequence and coordinates of the
159 primers are given in Table 3.2. For the synthesis/amplification of cDNAs
from internal portions of hrpS (Fragment A; Fig. 3.5) and hrpL (Fragment
D; Fig. 3.5), the primer pairs RTSR/RTSF and RTL/RTLF, respectively, were used. For the reverse transcription/amplification of hrpS-hrpY
(Fragment B; Fig. 3.5) and hrpX-hrpL (Fragment C; Fig. 3.5) inter-operon
cDNAs, primers RTYSR/RTYSF and RTLXR/RTLXF were used,
respectively. Per each 50-µl reaction, 2 µg of DNA-free total RNA was
mixed with 1 µM of each primer, 1X Titan RT buffer (with 1.5 mM
MgCl2 ), 0.2 mM dNTP mix, 5 mM DTT, 10 U RNase inhibitor
(Invitrogen), and 1 µl Titan enzyme mix (AMV and Expand High Fidelity;
Roche). Samples were incubated at 55°C for 30 min for reverse transcription, followed by PCR amplification. Initial denaturation was performed at 94°C for 2 min, followed by 30 cycles of denaturation at
94°C for 1 min, annealing at 55°C for 1 min, extension at 68°C for 2 min; final extension was at 68°C for 10 min. The “No RT” control was run following a standard PCR protocol using Taq DNA polymerase (Ausubel et al., 1987).
3.2.8 Northern hybridizations
For each strain, ca. 10 µg of total bacterial RNA was electrophoresed in a 0.9% agarose gel in 1X MOPS/formamide buffer as previously described (Ausubel et al., 1987). RNA gels were blotted overnight onto a
Nytran (S&S) nylon filter in 5X SSC buffer and fixed to the filter by UV
160 cross-linking following standard protocols (Sambrook et al., 1989). The
probes used to analyze the various transcripts were synthesized by PCR
using plasmid pMM58 as a template. Primers LFB and LRE (Merighi et
al., 2003) were used to make the probe for the hrpL transcript, primers X-
pMal-F and X-pMal-R for the hrpXY transcript and primers ET15SF and
ET15SR for the hrpS transcript. PCR products were labeled by random
priming using ([α32P] dATP and the Klenow fragment of DNA polymerase
I (Ausubel et al., 1987). About 2 x 107 cpm (Cerenkov) of probe were
used for each hybridization experiment. Prehybridization-hybridization
and post-hybridization wash conditions were performed at high stringency
conditions. The hybridization signals from the blots were detected by exposure to X-ray films (Kodak Biomax MS) at –80°C between two
intensifying screens for 24 to 48 h.
3.3 RESULTS AND DISCUSSION
3.3.1 Constitutive expression of HrpS indirectly upregulates hrpS
transcription by acting on the hrpL promoter
To confirm our preliminary results on hrpS-lacZ autoregulation, we analyzed gene expression in the strain MEX1 hrpS-lacZ carrying pRF205
+ Plac-hrpS after growth in IM5.5 for 16 h. (A map with relevant mutants
and plasmids is shown in Fig. 3.1.) β-galactosidase levels showed a modest but consistent upregulation of hrpS ranging from 1.5- to 2.5-fold
(Fig. 3.2) upon overexpression of hrpS from pRF205. A similar 161 experiment using strain DM733 hrpY::Tn5 hrpS-lacZ was performed. The results showed that the presence of the hrpY::Tn5 mutation strongly reduced expression of hrpS-lacZ in strain DM733 as compared to MEX1 and that the hrpS-lacZ fusion was not stimulated by overexpression of hrpS (Fig. 3.2). This finding is consistent with the model of regulation discussed in Chapter 2 and this latter result implies that HrpS is not acting directly on the hrpS promoter, but may require either HrpY or transcription from the hrpXY and/or hrpL promoters.
To verify whether the autoregulatory effect was due to transcriptional effects on the hrpXY promoter, we first assayed its expression in the presence and absence of pRF205 using strain MEX116 hrpX-lacZ as the genetic background. The results showed that hrpXY expression was upregulated ca. 4-fold by ectopic expression of HrpS (Fig. 3.2).
Upregulation of hrpX-lacZ by pRF205 was blocked in a strain carrying a second polar mutation in hrpL (strain DM2824 hrpL::Ω hrpX-lacZ) (Fig.
3.2), which was constructed by allele exchange of a hrpL::Ω gene into
MEX116 hrpX-lacZ. It is unlikely that HrpL is required for hrpXY upregulation by HrpS, because there is not a recognizable Hrp box in front of hrpXY, and the previous result suggests that the hrpXY promoter is not directly regulated by HrpS. Therefore, we hypothesized that the hrpL promoter may be required for the HrpS-dependent effect. This hypothesis was confirmed by assaying hrpS expression in strain DM2830
162 hrpL::Ω hrpS-lacZ in the presence and absence of pRF205 (Fig. 3.2).
Again, the autoregulation of hrpS observed in strain MEX1 hrpS-lacZ was
blocked by a polar mutation in hrpL.
Collectively these results suggest that HrpS increases hrpS-lacZ
expression via its action on the hrpL σ 54 promoter, which is part of the
linear cascade model described in Chapter 2. To explain how
autoregulation occurs, two possible scenarios can be hypothesized: (i) the
HrpS effect depends on increased expression of hrpXY via the hrpL σ 54 promoter, with subsequent activation of hrpS by HrpY as described in
Chapter 2 (“trans-effect”); or (ii) transcription from the hrpL σ 54 promoter could continue through hrpXY into hrpS (“cis-effect”). The first possibility is partially ruled out by the observation reported in Chapter 2
(Merighi et al., 2003) that a plasmid-borne hrpS-uidA gene fusion is not affected by hrpS mutations. In this experiment, we would have expected some downregulation of hrpS-uidA if the “trans-effect” hypothesis was correct, because reduced expression of hrpL, and consequently hrpXY, in the hrpS mutant would have ultimately reduced HrpY levels.
3.3.2 The upstream region of the hrpL promoter contains a symmetrical dyad required for HrpS-dependent activation
We wanted to test if the putative UAS site located at –163 from hrpL is required for the HrpS-dependent upregulation of this gene. The dyad-
163 symmetrical DNA element (TGCAA-N4 -TTGCA) weakly matches the E.
coli glnA enhancer site (Chapter 2). PCR fragments containing 134 and
208 bp upstream of hrpL were cloned into the GUS reporter plasmid pPL6GUSC to create plasmids pDM2785 and pMM191, respectively. The resulting plasmids were conjugated into strains DC283 and DM758 hrpS::Tn5 and then PhrpL-uidA expression was measured after growth in
IM5.5. The control reporter fusion in pDM2785, which contains all the
hrpL elements required for HrpS-dependent regulation, was highly
expressed in DC283 and reduced to basal levels in the hrpS background
(Fig. 3.3). However, the expression of the fusion in pMM191, which is
missing the putative UAS, was reduced to basal levels in both genetic
backgrounds (Fig. 3.3), thereby confirming that the deleted 74 bp region
contains an essential cis-acting sequence needed for normal regulation of
hrpL.
3.3.3 Primer extension analysis defines the 5’ ends of hrpXY, hrpS and
hrpL transcripts and their potential promoters
To further test the “cis-effect” model, we located the 5’ ends of the
hrpXY, hrpS and hrpL transcripts by primer extension analysis and
identified their potential promoters. As shown in Chapter 2, sequence
analysis revealed potential σ 70 promoters in front of hrpXY and hrpS and a
σ 54 promoter in front of hrpL. Primer extension analysis of mRNA
expressed under Hrp-inducing conditions showed that the hrpS mRNA 164 started 611 nt upstream of the ORF and included the entire abortive IS element (Fig. 3.4). Such a long untranslated leader sequence may form a strong (-149.2 kcal mol-1) hairpin structure, as discussed in Chapter 2.
This highly stable structure could complicate the analysis of transcripts by RT-PCR, reducing the efficiency of DNA polymerization or cause template switching. A putative σ 70 promoter is centered at –25 bp from the hrpS transcription start and has sequence (5’-GATGGTT-N17-TATTT-
3’). Primer extension of the hrpL transcript consistently produced a strong signal mapping at -39 nt from the hrpL start codon (Fig. 3.4). Visual
54 inspection identified a σ promoter (5’-CTGGCA-N6 -TTGCT-3’) centered at –19.5 bp from the transcript start. The hrpXY transcript start was mapped at –12 from the start codon, with a σ 70 consensus promoter
(5’-TGTGGA-N16-TTAA-3’) centered - 24.5 bp from the transcriptional start nucleotide(Fig. 3.4).
3.3.4 RT-PCR reveals that the hrpL-hrpXY and hrpXY-hrpS interoperon regions are transcribed
Oligonucleotide primers were designed to amplify cDNA transcripts spanning the hrpL-hrpXY and hrpY-hrpS intergenic regions and internal portions of hrpL and hrpS using a “one-tube” RT-PCR reaction (Fig.
3.5A; fragments C, B, D and A respectively). Total RNA was purified from P. stewartii DC283 grown in IM5.5 to early stationary phase. The control products from within the hrpL and hrpS ORFs (254 and 232 bp for 165 fragments A and D, respectively) were obtained, indicating that the hrp
regulatory genes were expressed under the experimental conditions used.
PCR products were also obtained using primers designed to amplify either the entire hrpXY upstream regulatory region (256 bp, fragment C; lane 2,
Fig. 3.5B) or the part of the hrpS regulatory region located upstream of the IS-like element (271 bp, fragment B; lane 4, Fig. 3.5B). Attempts to obtain PCR products with primers amplifying a product spanning the whole 5’hrpS region (fragment E; not shown in the gel figure) repeatedly failed, perhaps due to the strong secondary structure of the IS-like mRNA.
No product was amplified from the control without the reverse transcriptase enzyme, which was run using primers for the internal portion of hrpS and DNA-free total RNA as template (lane 6, Fig. 3.5B). These results indirectly show that no termination occurs downstream of hrpL and hrpXY.
3.3.5 Northern blot analysis reveals the presence of long transcripts spanning hrpL and hrpXY that are disrupted in a hrpL polar mutant
We attempted to confirm the results of the RT-PCR analysis by demonstrating the existence of long transcripts starting from PhrpL and spanning hrpXY and hrpS. We analyzed the transcripts of the hrp regulatory regions by Northern blotting using probes homologous to the hrpL, hrpX and hrpS genes (Fig. 3.6). DNA-free total RNA was prepared from strains DC283 (wild-type) and MEX105 hrpL::Tn3HoHoI grown in 166 IM5.5 to early stationary phase. Northern blot signals were generally
weak, probably due to the high-turnover rate of hrp messages, and
extended exposure for several days were required. In DC283, the hrpS
probe identified at least four transcripts of sizes 2.4 kb, 1.6 kb, 1.2 kb and
1.0 kb. The 2.4 kb and 1.6 kb signals were absent in the MEX105 hrpL::Tn3HoHoI strain, which still had the 1.2 and 1.0 kb transcripts
(Fig. 3.6). The negative control strain DM758 hrpS::Tn5 did not produce any signals with the hrpS or hrpL probes (not shown). The hrpL probe detected many transcripts in DC283 ranging in size from 2.9 kb to 0.7 kb
(Fig. 3.6), but only a 0.5 kb transcript was detected in strain MEX105 hrpL::Tn3HoHoI. Even if the long transcripts have a size shorter than expected, which could be due to rapid degradation of the mRNAs, these results are consistent with a regulatory model whereby transcription from the hrpL promoter continues into hrpXY and part of the hrpS regulatory region. This is consistent with the absence of clearly identifiable rho- independent terminators downstream of hrpL or hrpY. In contrast, potential terminators are present in the nucleotide sequence of this region from the related species E. amylovora, but a detailed transcriptional analysis has not been done in this bacterium. Therefore, this form of autoregulation may be a specific adaptation of P. stewartii.
167
3.4 CONCLUSIONS
The majority of the genetic and biochemical data presented in this
chapter support a model where hrpS is autoregulated at the transcriptional level via a DNA site located 134 bp upstream of hrpL, almost 3 kb from the hrpS 5’ regulatory region. This autoregulation does not appear to
depend upon trans direct activation of hrpS by either HrpS or HrpY,
because it was observed only when reporter fusions to hrpS were located in cis to the other regulatory genes. The effect of HrpS on its own
expression seems to be due instead to the production of a novel transcript
spanning the entire hrpL-hrpXY-hrpS region.
The results obtained from the RT-PCR analysis were consistent with the absence of transcriptional terminators downstream of hrpL and hrpY, thereby confirming the sequence analysis presented in Chapter 2. This result implies that interoperon transcription is possible in this system. We were not able to demonstrate the existence of the predicted 4.2 kb transcript originating from the hrpL promoter, but a 2.9 kb mRNA was identified in DC283 using the hrpL probe, which was absent in polar hrpL
mutants. Similarly, a 2.4 kb mRNA was detected in DC283 with the hrpS
probe and long transcripts were absent in the hrpL mutant. The estimated
sizes of the long transcripts in these experiments may not be accurate because our size standards did not contain fragments >3.5 kb. Also,
168 bacterial mRNAs undergo rapid turnover, so that the isolation of large,
undegraded transcripts is technically difficult. The model in Fig. 3.7
summarizes these findings.
Autoregulatory feedback loops are common in biological systems and
extensive literature on their mathematical modeling is available (Glass
and Kauffman, 1972; Thomas, 1973; Thomas, 1978). Most simple positive
feedback mechanisms lead to bistable systems, where two steady states
are observed. Modest perturbations of the system have little effect on the
level of expression, whereas major stimuli shift the system back and forth
between the two extreme levels of expression (e.g. on and off). The
regulatory loop triggered by HrpS appears to be a pure feedback
mechanism, at least at first approximation. If we consider the other
additional regulators entering the loop, the system may theoretically allow
for stable cycles or multiple steady states depending on whether negative
or positive regulators are involved in the feedback (Thomas, 1978). For
instance, additional known positive regulators are HrpY acting on hrpS
and HrpY acting on hrpXY. Negative regulators of hrpS are hypothesized
or shown to exist in Chapter 4 and Chapter 6. These negative regulators
may act by merely (re?)-inforcing the “off” state of the bi-stable switch more than being feedback controllers. Therefore the net effect is probably a system with at least two steady states.
Whether HrpY plays a direct role in reinforcing the HrpS-autoregulatory loop is not clear yet. The absence of any effect of hrpS mutations on a 169 plasmid-borne hrpS::uidA reporter (Table 2.5) suggests that changes in
the level of hrpXY expression arising from the feedback loop do not affect
hrpS significantly, possibly because the two-component system is already
at maximal expression under the growth conditions used. Therefore, one
may conclude that HrpY trans-activation of hrpS may not reinforce the loop significantly in the experimental conditions used in this study.
Alternatively, the fact that the hrpS-gusA fusion is plasmid-borne may alter the expected down shift in the hrpS mutant if one of the roles of
HrpY is to function as an antirepressor for hrpS transcriptional activation, as discussed in Chapter 4. In other words, a putative repressor of hrpS
may be titrated out by the plasmid-borne promoter fusion, therefore
masking the effect of HrpY.
An additional level of regulation of hrpS may involve a repressor of the interoperon transcription from hrpXY into hrpS. Indeed, hrpXY
transcription has a high, constitutive basal level in rich media.
Considering the absence of rho-independent terminators downstream of
hrpY, constitutive expression of hrpS would also be expected. However,
hrpS is repressed in rich media and only activated in planta or in IM5.5.
This issue is partially addressed in Chapter 4.
Determining whether the regulatory system works as an on/off switch or
as a rheostat with multiple levels of expression may be relevant to the
biology of the pathogen when it switches from living in the insect vector
to colonizing a corn leaf. From an experimental viewpoint, determining 170 this would require accurate quantification of the level of RNA and protein
expression over time and space using highly unstable reporters able to
quickly respond to the changing environment. The GUS reporters used in
this study are either not suitable for in planta expression analysis or are
too stable to indicate rapid decreases in expression. To this respect, the high turnover gfp[AAV] fusions described in Chapter 2 may offer new
tools to tackle these questions.
One possible biological role of the autoregulatory loop may be to
compensate for high turnover rates of the hrpS mRNA. Anecdotically, we had difficulties in amplifying by RT-PCR hrpS fragments from some RNA preparations, whereas the amplification of hrpL and hrpXY cDNAs was
always successful. This could reflect a greater instability for hrpS
transcripts. Measurements of turnover rates were not performed, so this
remains only a working hypothesis, but RNA stability has been shown to
play a role in regulation of hrp genes in different erwinias (Chatterjee et
al., 2002). As a negative note, an additional result previously published
by our lab (Frederick et al., 2001) seems to contradict the importance of
this autoregulatory loop during the infection of the plant. Indeed, a
Tn3HoHoI polar insertion between hrpL and hrpXY (strain MEX7) did not
affect virulence as measured by water-soaking production. This insertion
is expected to disrupt the autoregulatory loop at the transcriptional level.
One possible explanation is that under experimental infections, which
present less demanding and variable conditions than in natural field 171 infections, the autoregulatory loop may be optional. Future experiments should test whether virulence of MEX7 under field conditions is affected or not.
3.5 SUMMARY
Due to the particular arrangement of the hrpL, hrpXY and hrpS operons and the absence of clearly identifiable transcription terminators downstream of hrpL and hrpXY, activation of hrpL by HrpS can increase expression of HrpS leading to its apparent autoregulation. In this study, we used genetic and molecular biology approaches to demonstrate the existence of this autoregulatory loop and its dependence on a UAS located upstream of hrpL. The role for autoregulation in the disease cycle/biology of the disease and future studies are discussed.
172 Strain, or plasmid Relevant phenotype(s) and/or genotype(s) Source or reference
Escherichia coli
r q + + BW20339 F' 128::Tn10-12(Kan ) lacI ∆lacZM15 pro(BA) /DE3(lac)X74 uidA(∆MluI)::pir recA1
∆phoA532 ∆(phnC∆DEFGHIJKLMNOP)33-30 Metcalf et al. ( 1996)
DH10B F- mcrA ∆(mrr-hsdRMS-mcrBC) φ80dlacZ∆M15 ∆lacX74 endA1 recA1 deoR
∆(ara,leu)7697 araD139 galU galK nupG rpsL λ- Gibco BRL
- - - r HB101 F thi-1 hsd20 (r Bm B) sup E44 recA13 ara-14 leuB6 proA2 lacY1 rpsL20 (Sm )
xyl-5 mtl-1 Boyer and Iates Roulland
-Dussoix (1969)
HB101Rif spontaneous Rifr strain Lab collection
Sφ200Rif metB strA purB ∆(agg-uidA-man) Rifr Wei et al. (2000)b
P. stewarti subsp. stewartiii
SS104 Wild type, Wts+, HR+ ICPPBa
DC283 SS104 Nalr Coplin et al. (1986)
DM064 DC283 hrpY1296::Tn5 (Kanr) Frederick et al. (2001)
DM733 DC283 hrpY64::Tn5 hrpS1::Tn3HoHoI (Kanr, Ampr) Merighi et al. (2003)
DM758 DC283 hrpS224::Tn5 (Kanr) Frederick et al. (2001)
MEX1 DC283 hrpS1::Tn3HoHoI (Ampr) Frederick et al. (2001)
MEX116 DC283 hrpX116::Tn3HoHoI (Ampr) Frederick et al. (2001)
Plasmids pBluescript
KS and SK (+) ColE1 αlacZ (Apr) Stratagene pLAFR3 IncP cos αlacZ (Tcr) Staskawicz et al. (1987)
pRK415 IncP αlacZ (Tcr) Keen et al. (1988)
pHP45::Ω ColE1, (Apr) Frey and Krisch, 1985
pLD55 R6K ori, αlacZ, tetAR (Apr) Metcalf et al. (1996)
Table 3.1 (continued)
173 Table 3.1 (continued)
pPL6GUSC pLAFR6 derivative carrying a promoterless uidA gene (Tcr) Knoop et al. (1991)
+ - r r r pRK2013::Tn7 ColE1 mob traRK2 ∆ repRK2 repE kan::Tn7 (Tp , Sm , Sp ) Ditta et al. (1980)
pDM2533 pBSK carrying a 1.2 kb PCR fragment containing hrpL+ This study
pDM2862 pDM2533 with an Ω/SpR cassette at the NruI site of hrpL This study
pDM2868 pLD55 with the pDM2862 insert as XhoI/SstI fragment This study
pDM2785 pPL6GUSC with a 389 bp BamHI PCR fragment to create a hrpL-uidA fusion Merighi et al., (2003)
pDM1296 pES1044 hrpY1296::Tn5 (Kanr) Coplin et al., (1992)
pRF205 pVK100 with the 1.7 kb HindIII fragment containing hrpS from pES1044 Frederick et al. (1993)
pMM25 pPL6GUSC with a 244 bp HindIII-BamHI PCR fragment to create a
hrpX-uidA fusion Merighi et al. (2003) pMM50 pPL6GUSC with a 1.3 kb BamHI PCR fragment to create a hrpS-uidA fusion Merighi et al. (2003) pMM191 pPL6GUSC with a +220 to –95 hrpL PCR fragment fused to uidA This study
a International Collection of Plant Pathogenic Bacteria
Table 3.1. Bacterial strains and plasmids.
174 Primer Sequence (5'Æ3') Sequence coordinates (5') Source
RTSR4650 GTAAGCTAAGTTGTTGAC 4650-4633 This study
RTSF4397 ACTCATTTCGACCACATC 4397-4414 This study
RTLR931 GGGCAATACCAGATACCC 931-914 This study
RTLF700 AACCTGCACCCTACTGAG 700-683 This study
RTYSR3692 GCTGATATCGGGTAAGGA 3692-3675 This study
RTYSF3422 CATAAAGCCCGCCTGATG 3422-3439 This study
RTLXR1424 GATTGGCGTATCACTGCT 1424-1407 This study
RTLXF1169 CTGTTGGCACAGTACGTT 1169-1152 his study
LFB CGCGGATCCACATTAAGCCAAACGGCAAAA 662-642 Merighi et al., 2003
LRE CCGGAATTCTGTTTTACCCCGTTCAGTT 1255-1237 Merighi et al., 2003
XPMALR CGGAATTCTCACTCAGGGTATTAAAGGAATGGTC 2816-2837 This study
XPMALF GGGAATTCATGCAGTATTTGGACAGAC 1380-1398 This study
ET15SF CTACATATGAATATTGAAAATAATGAGCACTCATTCCGA 4354-4383 This study
ET15SR CGCGGATCCCAATCAGTCAGACAATGATGCGTTGCGTT 5301-5327 This study
LRBG CGCGGATCCTCTAGAATGACTTCCAGCCAGGTCAT 869-850 Merighi et al., 2003
LFU- CGCGGATCCGAGCTCAAAGTTGCGAAAAAATGAAGG 550-570 This study
PEX GCACGCTTCGTCATTGACAT 1498-1517 This study
PEL TATGTGGGGATGGAGGGTTAACGTG 724-748 This study
PES-1 CGATAGGTTGCTCTTTCGTGAATG 4412-4435 This study
PES-2 TCACCGACTCGCAATAATCC 4286-4305 This study
PES-3 TGGCTGGCTCACAATAACTG 4168-4187 This study
PES-4 TAGACAGGCAGCATGTGTTC 4024-4043 This study
PES-5 TGCGAGCCGCTACACTTATC 3839-3858 This study
PES-6 CGGGTAAGGAATTGATGGAG 3665-3684 This study
Table 3.2. Primers
175
hrpY1296::Tn5 hrpL2868::Ω
hrpS hrpY hrpX hrpL
MEX1 MEX116
HindIII HindIII XhoI SstI Plac pRF205 pDM2868
BamHI
pDM1296
Figure 3.1. Physical map of the hrp regulatory region, mutants and plasmids.
Open boxes represent open reading frames and the solid box represents the IS element remnant. Solid circles indicate Tn5 insertions, open circles indicate Ω interposon insertions, and arrows indicate Tn3HoHoI-lacZYA insertions. The inserts of the plasmids used in this study are shown beneath the map. The arrows also indicate the direction of transcription. MEX1 is DC283 hrpS-lacZ and MEX116 is
DC283 hrpX-lacZ.
176 3500 3000 2500
2000 none 1500 +pRF205 1000 500
B-Galacoside activity (pmol MU/min/OD600) 0
830 EX1 2 M DM733 M2824 M D MEX116 D
Figure 3.2. Effects of HrpS+ constitutive expression on chromosomal hrp-lacZ
fusions in various genetic backgrounds.
P. stewartii strains were grown in IM5.5 to OD600> 0.7 and ß-galactosidase assays
were performed as described in Materials and Methods. Enzyme units are pmol MU
per min per OD600. Strain MEX1 is DC283 hrpS-lacZ, strain DM733 is DC283 hrpY::Tn5 hrpS-lacZ, strain DM2824 is DC283 hrpL::Ω hrpS-lacZ, strain MEX116 is
DC283 hrpX-lacZ, and strain DM2830 is DC283 hrpL::Ω hrpX-lacZ.
177
54 DC283 DM758 σ IHF UAS hrpL hrpJ
5793±10 62±2 GUS pDM2785
71±2 54±1 GUS pMM191
100 bp
Figure 3.3. The effect of deleting the upstream enhancer sequence (UAS) in the hrpL regulatory region on PhrpL expression.
Expression of reporter plasmids pDM2785 and pMM191, carrying 208 bp and 134 bp, respectively, of the 5’ hrpL region fused to uidA, was analyzed in strains DC283 and DM758 after growth in IM5.5. pMM191 lacks the putative UAS. A map of the hrpL-hrpJ intergenic region and the inserts cloned into the reporter plasmids is shown. Data are means from two experiments with two replicates each. GUS activities
-1 –1 are reported as pmol MU min OD600 ± standard deviation.
178
TGCA T G C A 5’ T G C A 5’ C T C G G 5’ C +1’ T T A A T G T -10 G -12 A T C C T T C C A G T T G T C T A G T T C -10 A G C A C A C C T A C T A C A T T A G T G +1 G +1 T T C T A C A A A G A C +1 C A T A C T C G C T G A T C A C 3’ G G hrpL hrpX hrpS G 3’ 3’
Figure 3.4. Primer extension analysis of hrpL, hrpX and hrpS transcripts.
Total RNA from DC283 cells grown in IM5.5 was annealed with the 32P-end labeled primers PEL, PEX and PES-5 shown in Table 3.2 and extended as described in
Material and Methods. Lanes A, T, G, and C show the dideoxy sequencing ladders for
the hrpL, hrpX and hrpS genes generated with the same primers. The transcriptional
start sites are indicated by solid arrows. Mapping of the hrpXY 5’ mRNA end always
produced very weak signal (+1), with often a secondary signal locate 21 nt further
upstream (+1’). The lane corresponding to the hrpX cDNA was enhanced with
Photoshop (Adobe). The start codons (ATG) are highlighted. RBS = ribosomal binding
site.
179 A 254 bp 271 bp 256 bp 232 bp A B C D RTSR → RTYSR → ← RTYSF RTLXR → ← RTLF
← RTSF RTLXR → 1229 bp ← RTLXF E
hrpS hrpY hrpX hrpL
B 1 2 3 4 5 6 7
1 kb
0.5 kb
Figure 3.5. RT-PCR analysis of interoperon read-through transcription.
(A) The products amplified by RT-PCR and their sizes are shown as black bars above the genetic map (amplicons A, B, C, D, E). The black box upstream of hrpS represents the abortive IS element. The long transcript hypothesized in this study and mRNAs from each operon are shown below the map as arrows. (B) To demonstrate inter-operon transcription, total RNA was isolated from strain DC283 grown in IM5.5. mRNAs were reverse transcribed and cDNA was amplified using specific primer pairs to obtain the PCR products shown in the gel. Lane 1: 1 kb-plus molecular weight markers (Invitrogen); Lane 2: control without reverse transcriptase; lane 3: fragment
A; Lane 4: fragment B: lane 5: fragment C; lane 6: fragment D; lane 7: 1 kb-plus molecular weight markers.
180
Figure 3.6. Northern blot analysis of hrp regulatory transcripts.
(A) Northern blot autoradiographs. Total RNA was isolated from DC283 and
MEX105 hrpL::Tn3HoHoI grown in IM5.5. About 10 µg of RNA was separated in a formamide/MOPS gel and blotted on a nylon filter. The filter was then hybridized at high stringency with either a hrpS or a hrpL probe, labeled by random priming with
[α 32P]dATP. The washed filters were exposed to X-ray film for 48 hrs. Sizes of the transcripts in kb are indicated next to arrows and were determined using E. coli and
Arabidopsis thaliana rRNA as molecular mass standards. (B) Gene map with the probable position of the observed transcripts and the map position of the probes used
(other transcripts could by hypothesized if one considers hybridizations not spanning the whole probe). The symbol indicates a Tn3HoHoI transposon insertion.
181
182
HrpS
~P
hrpS hrpY hrpX hrpL
Activation Hrp regulon
Figure 3.7. Model of the hrp regulatory cascade including the autoregulatory loop
triggered by HrpS
This regulatory model is an extension of the linear model discussed in Chapter 2.
The activation of hrpL by HrpS is not only part of the basic cascade that activates the
Hrp regulon, but also triggers a positive autoregulatory loop that increases the levels
of hrpXY and hrpS transcripts. This loop may reinforce the autoregulatoy loop
triggered by HrpX/HrpY, thereby stabilizing the “on” state of the regulatory circuit.
183
CHAPTER 4
BIOCHEMICAL ANALYSIS OF THE RESPONSE REGULATOR
HRPY AND OF ITS ROLE AS A DUAL ACTIVATOR/REPRESSOR
OF HRP GENES
4.1 INTRODUCTION
hrp/hrc genes are required for infection and colonization of plants by necrogenic bacteria of the genera Pseudomonas, Xanthomonas, Erwinia
and Pantoea (Alfano and Collmer, 1997). They also play a role in rapid
establishment of soft rot erwinias, where enzymatic tissue disintegration
is only delayed in a hrp mutant (Rantakari et al., 2001; Perombelon,
2002). P. stewartii has a group I hrp gene cluster (Frederick et al., 2001), similar to those in Pseudomonas, Erwinia, Pectobacterium and other
Pantoea species, which is regulated by the HrpS enhancer and the ECF alternative sigma factor HrpL (Hutcheson, 1999). As shown for Pantoea agglomerans pv. gypsophilae and Erwinia amylovora, P. stewartii hrp genes are also regulated by the HrpX/Y two-component system (Wei et
al., 2000b; Merighi et al., 2003; Nizan-Koren et al., 2003). In all cases where mutational analysis has been performed, the HrpY response
184 regulator is essential for pathogenicity, whereas nonpolar mutations in the
hrpX sensor kinase gene have modest or no effects on pathogenicity (Wei
et al., 2000b; Merighi et al., 2003; Nizan-Koren et al., 2003). HrpY has
sequence similarity to response regulators of the FixJ class, which
includes E. coli NarL and UhpA and Bacillus subtilis DegU (Merighi et
al., 2003). Members of this class share a typical modular organization
(Stock et al., 2000), with a conserved N-terminal receiver domain (shared
by all classes of response regulators), a flexible linker, and a C-terminal
domain that contains a LuxR/GerR superfamily DNA-binding motif
(Heinikoff, 1995). The N-terminal domain of P. stewartii HrpY contains
three conserved aspartyl residues (D11, D12 and D57). By homology to
other response regulators, D57 is expected to be the phosphorylation site.
In agreement with this prediction, conservative and structurally neutral
amino acid substitutions at this position, such as D57N and D57A, abolish
the activity of hrpY when expressed in single copy. However, these
mutant proteins can fully complement hrpY null mutants for virulence or
HR elicitation when overexpressed from plasmids (Merighi et al., 2003).
Similar findings have been reported for E. coli UhpA (Kadner, 1983;
Kadner, 1995; Webber and Kadner, 1995). In all these cases, the
implication is that both the phosphorylation status of the response
regulator and its concentration in the cell are important parameters for its
activity and that the requirement for phosphorylation can be bypassed by
overexpressing it.
185 Using a genetic approach, we previously showed that P. stewartii HrpY
activates expression of hrpS and autoregulates the hrpXY operon (Merighi et al., 2003; and Chapter 2). This was confirmed by genetic reconstruction of each step in E. coli, which suggests that the effects of HrpY could be direct, unless HrpY is trans-regulating conserved factors in E. coli
(Merighi et al., 2003). However, the same set of genetic reconstruction experiments unexpectedly revealed that HrpY also represses the basal level of hrpL transcription when HrpS is absent. Inspection of the sequence of the regulatory regions of hrpS and hrpL did not reveal any apparent common sites that might explain the opposite effect of HrpY on these two promoters, but the regulatory sites bound by FixJ-class regulators are often quite degenerate (Tyson et al., 1994; Dahl et al.,
1997).
Although extensive genetic data is available for the regulatory role of
HrpY in P. stewartii, P. agglomerans pv. gypsophiliae and E. amylovora, a biochemical analysis of the target regulatory regions has not been done and it is not known how HrpY activates transcription. In this study, we define the portions of the hrpS and hrpL 5’ regions required for HrpY- dependent regulation and we provide biochemical evidence for the interaction of recombinant HrpY with defined regions of the two hrp promoters. We also show that HrpY has a single phosphorylation site located at D57 and that its phosphorylation increases the affinity of HrpY for the DNA sites in front of the hrpS promoter. Specific binding of HrpY
186 to the hrpL promoter at low affinity was also demonstrated in vitro.
Preliminary DNase I footprints for the hrpS regulatory region and a
working model explaining most of our genetic and biochemical data are presented.
4.2 MATERIALS AND METHODS
4.2.1 Strains and growth conditions
E. coli and P. stewartii strains and plasmids used in this study are listed in Table 4.1. Luria-Bertani (LB) broth and agar were used for strain
maintenance and clonings. To induce hrp genes in P. stewartii strains, we
followed the procedure described in Chapter 2 using the IM pH5.5
(IM5.5) liquid medium. Liquid cultures were grown in flasks or tubes with
shaking at 200 rpm at 37°C for E. coli or 29°C for P. stewartii. Special
growth conditions were used for growing E. coli BL21(DE3) strains
during expression purification of various recombinant proteins, as
described below. When appropriate, antibiotics were supplied at the
following concentrations: ampicillin, 200 µg ml-1; kanamycin, 50 µg ml-1; and tetracycline, 20 µg ml-1; chloramphenicol, 34 µg ml-1.
187 4.2.2 Construction of plasmid-borne gene fusions and of a hrpY[D57E] allele for plasmid expression
Unless otherwise stated, isolation and manipulation of recombinant
DNA molecules used standard procedures (Ausubel et al., 1987) or the various product manufacturers’ instructions. Fragments containing deletions of the 5’ regulatory region of hrpS were produced by PCR using the primers given in Table 4.2. Oligonucleotide primers were synthesized at IDT-DNA (Iowa). All PCR products were generated with 15 to 20 cycles of amplification using Pfu DNA polymerase (Clontech) and 500 ng of plasmid pMM58 as a template. Gel-purified products were digested with BamHI and ligated into pPL6GUSC to create transcriptional fusions to a uidA (GUS) reporter gene (Fig. 4.1). In particular, the hrpS fragment from–231 to +628 was amplified with primers SF3509B and SR4368B and the corresponding reporter plasmid was named pMM400. Region –165 to
+628 was amplified with primers SF3575B and SR4368B and the reporter plasmid constructed was named pMM396. Region –90 to +628 was amplified with primers SF3650B and SR4368B and the corresponding plasmid was named pMM393. Region –51 to +628 was amplified with primers SF3689B and SR4368B and the corresponding plasmid was named pMM403. Finally, region –5 to +628 was amplified with primers SF3735B and SR4368B and the corresponding plasmid was named pMM392.
Construction of the two reporter plasmids carrying different fragments of the hrpL regulatory region is described in Chapters 2 and 3. The insert
188 in plasmid pDM2785 spans from –148 to +220 of hrpL (with +1 being the
transcription start at the σ 54 promoter identified in Chapter 3) and plasmid
pMM191 carries the –94 to +220 fragment. A third hrpL reporter gene
54 fusion lacking sequences downstream from the σ -promoter region
coordinates –148 to –12) was constructed by PCR, using primers JFGB
(a.k.a. HRPLP1) and L637R. The resulting plasmid was named pMM428
(Fig. 4.2).
4.2.3 Construction of plasmids for protein expression and purification
PCR was performed using Pfu DNA polymerase (Clontech), primers ET-
YF-NdeI and ET-YR-NdeI (Table 4.2), and plasmids pMM46 hrpY+ or
pMM94 hrpY[D57N] as DNA templates (Merighi et al., 2003). Primer ET-
YF-NdeI introduced an NdeI site at the start codon of hrpY, which maintains the reading frame with the vector’s N-terminal His6 -tag and
primer ET-YR-NdeI introduced three stop codons downstream of the hrpY
ORF. The ca. 670-bp PCR fragment, spanning hrpY+ or hrpY[D57N], was
gel-purified, digested with NdeI, and ligated into an NdeI-linearized pET-
15b plasmid (Novagen). The resulting plasmids, pMM221 and pMM222,
carried hrpY and hrpY[D57N], respectively, fused to an N-terminal His6 -
tag and expressed from a T7lac inducible promoter. The inserts in
plasmids pMM221 and pMM222 were sequenced using a cycle sequencing
protocol with fluorescent terminators and an ABI3700 capillary sequencer
at the OSU Plant-Microbe Genomics Facility. 189 4.2.4 Purification of His6 -HrpY proteins
Plasmids pMM221 and pMM222 were transformed by electroporation
into E. coli BL21(DE3) carrying pLysS. Several independent Apr Cmr isolates were stored at –20 and -80°C in 40% glycerol, 10 mM MgSO4 , 50 mM Tris-HCl pH 7. For protein expression, bacteria were grown overnight in 5 ml LB broth with antibiotic selection. A 3-l flask containing 1000 ml
LB broth was inoculated with 2.5 ml of overnight culture. The flask was shaken at 37°C until the culture reached A600=0.8 (measured in a B&L
Spectronic 20 colorimeter with 13 mm x 100 mm tubes). IPTG was added
to a final concentration of 1 mM and the culture was grown at 15°C for 18
to 19 h until A600=1.4. Cells were harvested by centrifugation in a Sorval
GSA rotor at 8000 rpm for 20 min at 4°C and then resuspended in 35 ml
of ice-cold lysis buffer (10 mM imidazole, 500 mM NaCl, 1 mM MgCl2 , 1 mM PMSF, 20 mM Tris-HCl pH 7.9 at 4°C), containing 50 µl of protease inhibitor cocktail (Sigma Cat. P8849). The cell suspension was incubated in 0.1 mg ml-1 lysozyme for 20 min on ice and then passed two times
through a 40-K French pressure cell (Thermo IEC) at 10,000 psi. Ten 5-s
bursts were applied with a Biologics Model 150V/T sonicator at 20% duty
cycle to further reduce the viscosity. The lysate was centrifuged twice at
20,000 x g for 15 min at 4°C. . Column chromatography was performed at
4°C. The supernatant was then applied to a 1.5 ml Ni2+-IDA column (I.D.=
1 cm; Sigma Cat. 56000) packed using His-Bind™ resin (Novagen),
190 previously loaded with 50 mM NiS04 and equilibrated with lysis buffer.
The column was washed with ten column-volumes of lysis buffer and ten
column-volumes of washing buffer (40 mM imidazole, 500 mM NaCl, 1
mM MgCl2 , 0.5 mM PMSF, 20 mM Tris-HCl pH 7.9 at 4°C). The target
His6 -HrpY proteins were eluted with 6 ml each of 100 mM and 250 mM
imidazole buffers (100 mM or 250 mM imidazole, 500 mM NaCl, 20 mM
Tris-HCl pH 7.9 at 4°C, 1 mM MgCl2 , 0.5 mM PMSF) in 1 ml fractions.
Fractions containing higher concentrations of the 26.1 kDa recombinant
protein, as assayed by SDS-PAGE, were pooled and dialyzed overnight
against 2 l of storage buffer (67 mM potassium glutamate, 125 mM
HEPES pH 7.9 at 4°C, 0.5 mM DTT, 0.5 mM PMSF, 20% glycerol). The
semipurified protein preparation was stored at –80°C until the next step.
The protein was further purified by anion exchange chromatography.
Preparations were diluted 1:10 in buffer A (20 mM Tris-HCl pH8.0 at
4°C, 25 mM NaCl, 0.25 mM EDTA, 0.2 mM PMSF), dialyzed twice for 2
h against 1 l of buffer A and loaded onto a 5-ml Q-Sepharose (Amersham-
Pharmacia) column prepared according to the manufacturer’s instructions.
The column was washed with 15 ml buffer A, proteins were eluted in a
linear buffer gradient prepared by mixing 30 ml buffer A with 30 ml
buffer B (20 mM Tris-HCl pH8.0 at 4°C, 500 mM NaCl, 0.25 mM EDTA,
0.2 mM PMSF) using a 100-ml gradient former (GIBCO) and 1 ml
fractions were collected.. Fractions containing highly purified HrpY were pooled and the buffer was exchanged for 1X storage buffer (10 mM Tris- 191 HCl pH 7.9, 125 mM NaCl, 1 mM DTT, 3 mM MgCl2 , 38% glycerol) by
repeated centrifugation in a Vivaspin 15R ultrafiltration tube (Viva
Science) with a molecular weight cut-off of 10 kDa following the
manufacturer’s instructions. Protein concentration was determined by the
dye-binding method of Bradford (Bradford, 1976). The degree of
purification was estimated by Coomassie Brilliant Blue staining
(Laemmli, 1970).
4.2.5 Mass spectrometry
Protein molecular weight measurements were performed by electrospray
ionization mass spectrometry using a Micromass Q-TOFTM II (Micromass,
Wythenshawe, UK) mass spectrometer at the OSU CCIC-MS Facility.
About 300 pmol of protein in storage buffer was concentrated and desalted in C18-solid phase manual syringe traps (Michrom BioResources) following the manufacturer’s instructions and then eluted in 50% acetonitrile, 0.1% TFA. Protein samples were introduced by infusion at 10
µl/min. Electrospray ionization was performed with a capillary voltage of
3.5 kV, source temperature of 110°C, cone voltage of 60 V, and a coaxial nitrogen flow. The quadrupole 1 range was set from m/z 100-1800 and all transmitted ions were scanned over m/z 100-3000. Data were acquired in continuum mode for 2 to 5 min. m/z spectral data were deconvoluted in the 22 to 30 kDa region using MaxEnt software (Micromass).
192 4.2.6 In vitro phosphorylation of HrpY
HrpY was phosphorylated using two alternative methods. To perform
chemical phosphorylation, a mixture containing 110 µM HrpY and 50 mM
carbamoyl phosphate (Sigma Cat. C5625) in phosphorylation buffer (125
mM potassium glutamate, 125 mM HEPES pH 8.0, 5 mM MgCl2 , 0.1 mM
DTT, 5% glycerol) was incubated at 25°C. For gel shift assays,
phosphorylated HrpY preparations were serially diluted in
phosphorylation buffer containing 50 mM NaCl. For enzymatic
phosphorylation, 15 µl of a mixture containing 25 µM HrpY, 75 µM
Salmonella enterica BarA198 (Teplitski et al., in press), 40 µM [γ32P]ATP
(10 Ci/mmol; NEN) and 0.1 mM cold ATP in phosphorylation buffer was incubated at 25°C for 90 min. Reactions were terminated with 5 µl of 4X stop buffer (125 mM Tris-HCl pH 6.8, 8 mM EDTA, 4% SDS, 8% β- mercapto-ethanol, 20% glycerol, 0.02% bromophenol blue), separated by
SDS-PAGE (12% monomer gel), and visualized by by Coomassie Brilliant
Blue staining (Laemmli, 1970). Gels were dried under vacuum at 60°C for
30 min. The radiolabeled proteins were visualized with a phosphor storage screen, analyzed on a Molecular Dynamics PhosphorImager and quantified with ImageQuant software.
193 4.2.7 Electrophoretic mobility shift assay
DNA fragments used as probes for electrophoretic mobility shift assays
were made by PCR using plasmid pMM58 (Merighi et al., 2003) as
template. Six probes (A, B, C, C1, C2 and C3) were designed to amplify
various fragments of the hrpS regulatory region located upstream of the
IS-like element. Primers SF3509 and SR3845 were used for amplification
of fragment A (327 bp); SF3509 and SR3575 for fragment B (76 bp);
SF3575 and SR3645 for fragment C (261 bp); SF3575 and SR3650 for
fragment C1 (76 bp); SF3650 and SR3689 for fragment C2 (86 bp); and
SF3689 and SR3845 for fragment C3 (101 bp). Six probes were also
synthesized to span the hrpL regulatory region (F, G, H, I, L and M). PCR
reactions used the following primer pairs: LRBG and L638F for probe F
(232 bp); LRBG and L620F for probe G (250 bp); LRBG and L578F for
probe H (292 bp); LRBG and L555 for probe I (315 bp); LRBG and JFBG
for probe L (389 bp); and L638R and JFBG for probe M (156 bp). PCR
products were purified from agarose or acrylamide gels and quantified by
the ethidium bromide staining spot method (Ausubel, 1997). DNA
fragments were labeled at the 5’ ends by incubation with T4
polynucleotide kinase (USB) and [γ32P]ATP (3000 Ci/mmol; NEN).
Unincorporated nucleotides were removed by Sephadex G-25 spin
chromatography (Sambrook et al., 1989) and labeled DNA was diluted to
10,000 cpm µl -1 (Cerenkov counts). DNA binding reactions (20 µl) contained 15 to 50 fmol 32P-labeled DNA probe, different amounts of 194 HrpY, 200 ng acetylated BSA (NEB), 1 µg poly(dI-dC) (Roche) in binding
buffer (125 mM potassium glutamate, 125 mM HEPES pH 7.9, 75 mM
NaCl, 5 mM MgCl2 , 0.1 mM EDTA, 0.2 mM DTT). Unlabeled specific
competitor DNA was added before the addition of HrpY. The reactions
were incubated at 25°C for 30-40 min. Reaction mixtures were loaded,
without loading buffer, onto a 1.5 mm-thick 6% acrylamide nondenaturing
gels (19:1 mono:bis ratio) in a V16 vertical gel apparatus (Waterman), run
into the gel at 25 V, and then separated at 200 V for 2-2.5 h. Gel and
running buffers were 1X TBE (97 mM Tris-HCl, 90 mM borate, 3 mM
EDTA, pH 8.0). The gels were dried under vacuum at 80°C and the
radioactive fragments were visualized either by autoradiography at –80°C
with Kodak MS films or on a phosphor storage screen and scanned on a
Molecular Dynamics PhosphorImager for quantification.
4.2.8 DNase I footprinting
Primer SF3509 was end-labeled with [γ32P]ATP (3000 Ci/mmol; NEN) using T4 polynucleotide kinase (USB) and unincorporated nucleotides were removed by Sephadex G25 spin chromatography. HrpS promoter fragments basically corresponding to probe A were amplified with SF3423 and SR3835 using pMM58 as a template. PCR products were purified from a 1% low-melting point agarose gel and recovered DNA was resuspended in TE buffer. DNA-protein binding reactions were performed as described for gel-shift assays using increasing concentrations of HrpY and 10 ng of 195 hrpS promoter DNA. All reactions contained 5 mM CaCl2 . The equilibrated complexes were subjected to partial digestion with protease- free DNase I (Worthington Biochemicals Cat. LS006342) diluted to 10
-1 units ml in binding buffer with 0.01% NP40 and 40 mM CaCl2 .
Digestion was performed at 25°C for 2 min and terminated with 3.5-vol stop buffer (400 mM sodium acetate pH 5.5, 0.2% SDS, 10 mM EDTA, 50
µl ml-1 yeast tRNA) and DNA was precipitated by adding 2.5 vol 100%
ethanol. Partially digested complexes were amplified by linear PCR in a
15 µL reaction mixture containing 1X PCR reaction buffer (Invitrogen), 2
32 mM MgCl2 , 2 pmol P-labeled SF3423 primer, and 0.2 U Taq DNA
polymerase (Invitrogen). The amplification cycle consisted of
denaturation at 95°C for 2 min, followed by 30 cycles of denaturation at
95°C for 30 s, annealing at 50°C for 30 s, extension at 72°C for 30 s.
Labeled DNA was separated on 6 % polyacrylamide gels in 1X TBE buffer with 8 M urea. A and G sequencing ladders were produced from fragment
A by PCR cycle sequencing using dideoxyterminators from the fmolTM
Cycle sequencing kit (Promega) and run alongside the DNase I digestion patterns. Gels were dried under vacuum at 80°C and exposed to X-ray films at –80°C for several days.
196 4.2.9 β-glucuronidase enzyme assays
β-glucuronidase (GUS) activity was assayed fluorometrically using 4-
methyl-umbelliferyl-β-D-glucuronide as described by Jefferson (1987),
but the assays were scaled down to fit microtiter plates and analyzed
using a Victor 1420-2 multilabel reader (PE-Applied Biosystems). Net
GUS activity of each strain was corrected for the basal fluorescence of P.
stewartii DC283 or E. coli SΦ200 carrying pPL6GUSC without an insert.
-1 -1 - Specific activity was expressed in GUS units (pmol MU min OD600 ml
1 of culture at 37°C).
4.2.10 Total RNA isolation
Total bacterial RNA was isolated from 5 ml of overnight cultures of
cells grown in IM pH5.5 liquid medium or 5 ml of exponentially growing
cultures in LB broth. Cells were pelleted and resuspended in 0.1 ml of
TEN100 buffer (10 mM Tris-HCl, 100 mM NaCl, 1 mM EDTA, pH 7.0).
Five volumes of Trizol (Invitrogen) at 65°C was added to the resuspended
cells, which were then vortexed, shaken at 65°C for 10 min and
centrifuged at 10,000 x g for 30 min at 4°C. The upper aqueous phase was
extracted with 0.1 ml of water-saturated chloroform and the RNA was
precipitated by adding one volume of isopropanol and centrifuging as
above. RNA was resuspended in 50 µl of DEPC-treated water and treated
with DNase I (Invitrogen) according to the manufacturer’s instructions.
197 4.2.11 Primer extension and DNA sequencing
Analysis of the 5’-ends of transcripts was performed by primer
extension (Sambrook et al., 1989). Primer PEL (Table 4.2) was labeled
with [γ32P]ATP and cDNA synthesis was carried out using M-MLV reverse transcriptase (Promega) with 50 µg of RNA per reaction following
the manufacturer’s instructions. DNA sequencing was performed with the
same primers used for cDNA synthesis following the fmolTM cycle sequencing protocol from Promega. The labeled extension products were visualized by autoradiography with Kodak XOMAT films after electrophoresis in 6% denaturing acrylamide gels at 55 Watts in a Bio-Rad sequencing apparatus (50 x 22 cm gel plate). For weakly labelled cDNAs, a second autoradiography exposure was performed, and the DNA ladder and cDNAs were merged electronically using Photoshop Elements
(Adobe).
4.2.12 Sequence analysis and modeling
DNA and protein sequences were aligned using ClustalX. Modeling of the HrpY tertiary structure was performed using a homology approach with the Swiss-Model algorithm at www.expasy.ch. The model was
checked for structural errors with the protein verification tool
WhatCheck. DNA curvature propensity and bendability was calculated
using Bend-It at www.icgeb.trieste.it.
198 4.3 RESULTS
4.3.1 Deletion analysis of the regulatory elements activated or
repressed by HrpY
To identify the sequences required for activation of hrpS and
repression of hrpL by HrpY, we constructed nested deletions of the
corresponding 5’-regulatory regions of each gene by PCR and fused the
resulting promoter fragments to the reporter gene uidA. The region
between hrpY and hrpS spans 844 nucleotides and includes a 483-bp-long
IS-like element located at –23 bp from the hrpS start codon. Primer
extension experiments described in Chapter 3 revealed a transcript
starting 106 nt from the 5’ end of the IS element (Fig. 3.4) with a σ 70 promoter nearby. We therefore focused our attention on only the region upstream of the IS-like element.
The DNA sequence of the region located between the end of hrpY and the start of the IS-like element (region +95 to –231 from the transcription start) in P. stewartii was aligned with the corresponding regions from P. agglomerans pv. gypsophilae, E. amylovora and Pectobacterium chrysanthemi to identify conserved and divergent sequence motifs. Cluster analysis grouped the sequences according to phylogenetic relationships
(not shown), with highest similarity between P. stewartii and P. agglomerans. Several blocks of highly conserved sequences were found in the most 5’ part of the region (here referred as CUR, conserved upstream region), followed by a more divergent short region (VUR = variable
199 upstream region) and then a putative IHF site (Fig. 4.3). Downstream of
the IHF site, two direct repeats were conserved among pantoeas (PDR =
Pantoea direct repeats). These were followed by a putative, highly
conserved σ 70-promoter, centered at –24 bp from the mapped transcription
start (Fig. 4.3). The curvature propensity and bendability of the
P.stewartii hrpS promoter region was also analyzed using the Bend-It algorithm (Fig. 4.4). Two peaks of potential intrinsic curvature were found around positions –129 and -147.
We tested in first approximation the function of these hypothetical elements by deletion analysis and construction of the reporter plasmids
described in Fig. 4.1. The plasmids were tested in wild-type and
hrpY[D57N] genetic backgrounds after growth in IM5.5. Plasmids
containing the entire hrpS 5’ regulatory region, such as pMM50 and
pMM400 (Fig. 4.1), required HrpY for activation. Removal of the region
from –165 to –231 (comprising the CUR) totally abolished hrpS
expression in both genetic backgrounds. Further deletion of the 5’ regions comprising the VUR (plasmid pMM393; Fig. 4.1) restored some
measurable basal activity, but the gene fusion showed much lower activity
compared to pMM400 or pMM50, and the expression was HrpY-
independent. A similar result was observed for pMM403, which carries a
further deletion spanning the two PDRs (Fig. 4.1) but contained the σ 70-
promoter. Plasmid pMM392 did not contain the σ 70-promoter and showed
even lower basal expression levels. 200 A detailed sequence analysis of the hrpL promoter region of P. stewartii is described in Chapter 2. Two hrpL reporter plasmids, pMM191 and pDM2785 were constructed for the studies described in Chapter 2 and
3. pMM191 lacks the UAS element required for activation by HrpS (see
Chapter 3). The plasmids were moved into E. coli SΦ200, the strains were grown in a rich medium and then the fusions were tested for a response to pMM46, which expresses hrpY+. The results shown in Fig. 4.2 confirmed the ability of hrpY to repress the full-length hrpL promoter (plasmid pDM2785; see also Chapter 2). A similar result was obtained with pMM191, which contains only the region up to –95 bp, without the UAS.
This implies that HrpY does not repress hrpL basal expression by interfering with the binding of HrpS (in P. stewartii) or its paralog NtrC
(in the E. coli SΦ200 reconstruction system) to the conserved UAS. While performing the primer extension experiments described in Chapter 3, we observed the presence of a second transcript starting at –58±1 from the
σ 54-promoter transcription start (Fig. 4.5). This transcript was not consistently found in all P. stewartii RNA preparations, but it was made in rich media. This implies the existence of a tandem promoter upstream of the σ 54-box. Sequence analysis using the NNPP algorithm indeed identified a putative σ 70 promoter centered at –68 (Fig. 4.5). We hypothesized that this promoter was responsible for the basal levels of hrpL expression and that it was repressed by HrpY. A third reporter plasmid, pMM428, containing the UAS element, but lacking the region 201 downstream of the σ 54-promoter, was constructed (Fig. 4.2). This hrpL- uidA fusion had a lower basal expression compared to pDM2785, and was no longer repressed by HrpY. Instead, a reproducible increase in hrpL expression was observed (Fig. 4.2). These results imply that sequences important for the repression of PhrpL by HrpY reside downstream of base –
12.
4.3.2 Overexpression and purification of HrpY proteins
HrpY and HrpY[D57N] were overexpressed using the pET-15b
expression plasmid. Inserts, produced by PCR, were engineered to maintain the correct translation frame with a vector-encoded N-terminal polyhistidine tag (ca. 2 kDa in size). To show that the his6 tag did not
affect biological activity, the his6 -hrpY gene was subcloned into pRK415.
Although this plasmid had an adverse effect on the growth of P. stewartii transconjugants, it weakly complemented a hrpY null mutant (data not shown). We decided to use the recombinant form of the protein for further studies due to the convenience of purification. Induction of the T7lac promoter of plasmids pMM221 (hrpY) and pMM222 (hrpY[D57N]) in E. coli BL21(DE3)(pLysS) by IPTG resulted in the production of 26-kDa polypeptides, which were not present in uninduced cells. (For HrpY see
Fig. 4.6A, lane 2 vs. 1; HrpY[D57N] is not shown). Fig. 4.6 (A to C) shows recovery of His6 -HrpY after various steps in its purification by
immobilized metal affinity chromatography using step-elution gradients
202 (Fig. 4.6 B) and subsequent anion-exchange chromatography of the
semipurified preparation in a linear salt gradient. Highly purified His6 -
HrpY was recovered from the 136 to 250 mM NaCl fractions (Fig. 4.6 C).
In some cases, ion-exchange chromatography was not necessary to obtain
highly purified HrpY, especially if washing buffers with higher concentrations of imidazole (60 to 80 mM) were used before the final elution step (100 to 150 mM). In general, the purity of the samples was assessed by densitometric analysis of Coomassie-stained SDS-PAGE gels.
Samples were also tested for the absence of background kinase activity using [γ32P]ATP. In all cases purity was > 95% and kinase activity was
not visible after autoradiography for 24 h. The average yield was of 8 mg
of purified protein per liter of culture. The protein stocks had
concentrations ranging from 150 to 350 µM in storage buffer and were
kept at –20°C until use.
The molecular weight of the polypeptides was confirmed by Q-TOF
electro-spray ionization mass spectrometry (Fig. 4.7). For the HrpY
sample, the major protein species had a molecular weight of 26051 ± 58
Da (Fig. 4.7A-B), whereas HrpY[D57N] was 26122 ± 76 Da (Fig. 4.7C-
D), both in close agreement with the expected mass of 26.1 kDa. Finally,
western blot analysis using anti-His6 monoclonal antibody confirmed the
identity of the recombinant polypeptides (not shown).
203 4.3.3 Phosphorylation of HrpY at D57 by Salmonella enterica BarA198
Response regulators are usually phosphorylated at a single Asp residue in the receiver domain, even if secondary phosphorylation sites are sometimes present (Delgado et al., 1993). The tertiary structure of HrpY was modeled using a homology approach based on the experimental crystallographic data from NarL and Spo0F (Fig. 4.8A) crystals or solution structures as templates (Protein Data Base files 1404.pdb,
1RNL.pdb, 1FSP.pdb, 1NAT.pdb). An E-score of 0.0001 was needed for the ProMod II algorithm to construct an approximated model of HrpY.
The model was energy-minimized with GroMol96 (Structure free energy =
-5742.091 KJ/mol) and the structure was checked with the What-If algorithm for detecting major structural problems. Modeling was successful from residue 3 to 206 out of the 216 making up the HrpY polypeptide. Several side-chain clashes and improper dihedral angles were found, but they did not involve the catalytic site (data not shown). The model predicts that three conserved aspartyl residues (D11, D12 and D57) at the top of a β–strand form the conserved catalytic triad (Fig. 4.8B).
Mutagenesis at D57 was simulated with SpdbViewer v. 7.0. Substitution of D57 with N, A or E and selection of the most energetically favorable rotamer did not cause clashes with nearby residues, implying that major structural changes are not expected upon mutagenesis (not shown).
We decided to directly test if D57 was the sole phosphorylation site in the response regulator. The putative cognate sensor kinase for HrpY is
204 encoded by HrpX because they are in the same operon and null mutations
in HrpX alter the Hrp phenotype (as shown in Chapter 2). Attempts to
purify sufficient amounts of active full-length HrpX for reproducible in
vitro trans-phosphorylation assays repeatedly failed (See Chapter 5).
Instead we used recombinant Salmonella enterica BarA198 (a truncated
version of BarA engineered with a N-terminal polyHis tag) for our
experiments. We selected this kinase because it was available from B.
Ahmer lab, a BarA ortholog (GacS) is present in all erwinias, and SirA
(the cognate response regulator for BarA) belongs to the same class of
regulators (FixJ) as HrpY. Fig. 4.9 shows that HrpY alone did not
catalyze an autophosphorylation reaction in the presence of [γ32P]ATP. On
the other hand, BarA198 autophosphorylated in presence of [γ32P]ATP
(Fig. 4.9, Lane 2) and phospho~BarA198 efficiently transferred 32P to
HrpY (Lane 3). The HrpY/BarA198 complex appeared more active than
BarA198 alone in catalyzing the autophosphorylation reaction, at least under the conditions used here (molar ratio kinase:response regulator monomers = ca. 3:1). Finally, the HrpY[D57N] mutant protein was not phosphorylated in presence of BarA198 and [γ32P]ATP (Lane 4),
demonstrating that D57 is the only phosphorylation site in HrpY.
Complete kinetic studies of the phosphotransfer and dephosphorylation
reaction were not conducted because we do not know whether or not the
205 use of BarA is a good model for the interaction of HrpX with HrpY. More information on Salmonella BarA biochemistry was recently published
(Teplitski et al., 2003)
4.3.4 Binding of HrpY to 5’ hrpS DNA fragments
The binding of HrpY to various DNA fragments spanning the hrpS regulatory region located between the IS-like element and hrpY was studied by electrophoretic mobility shift assays. Again, we focused on this region because primer extension and genetic experiments indicate that it contains the hrpS promoter. The fragments were synthesized by PCR and end-labeled with 32P. Their map positions are shown in Fig. 4.10.
Increasing concentrations of unphosphorylated HrpY (≤625 nM) were incubated with 25 fmol 32P-labeled fragment A (327 bp) for 40 min and then the protein-DNA complexes were separated from the free probe by electrophoresis in native 6% acrylamide gels (Fig. 4.11). A peculiar shift pattern was observed, with complexes of intermediate mobility migrating between the free probe and the samples with maximal concentration of
HrpY. The intermediate complexes formed wavy bands and may have been produced upon dissociation of the binding complex at lower HrpY concentrations. This has been reported in other systems (Dahl et al.,
1997). A similar result was obtained with the HrpY[D57N] variant (data not shown). All the reactions were performed in a large molar excess of the aspecific competitor poly(dI:dC). In a specificity control, binding of
206 HrpY to the hrpS fragment (A probe) was dramatically reduced by competition with a 100-fold molar excess of cold fragment A (Fig. 4.11).
These results demonstrate specific binding of HrpY to the full-length hrpS
promoter region.
To identify the HrpY-binding sites in fragment A (coordinates +95 to –
231), PCR probes carrying different portions of the hrpS promoter region
were synthesized and end-labeled with 32P. Binding assays were
performed with: probe B (-165 to –231) containing the CUR; probe C1 (-
90 to 165) carrying the CUR; probe C2 (-5 to –90) containing the DRs and
the σ 70-promoter; probe C3 (+95 to –5) containing the 5’UTR upstream of the IS-like element; and probe C (+95 to –165) that spanned C1, C2 and
C3 (Fig. 4.10). In each assay, 10-15 fmol of probe was incubated with 5
µM of HrpY with or without cold specific competitor DNA. HrpY formed high affinity complexes with fragment C, similar to what observed for probe A (Fig. 4.12). Significant amounts of fragment C2 were also retarded by HrpY and three complexes were observed. In a few experiments, fragment B was weakly bound by HrpY, but the sites must have had low affinity, because this binding was eliminated by competition with cold fragment C. Fragment C1 did not bind HrpY, whereas occasional binding was shown for probe C3.
To demonstrate the specificity of the binding of HrpY to fragment C, excess unlabeled fragments B, C1, C2 and C3 were incubated with HrpY and 32P-labeled probe C (Fig. 4.13). Up to 250-fold molar excess of 207 unlabeled fragments B, C1 and C3 did not block the binding of HrpY to
probe C, whereas binding was reduced in the control assay using cold
fragment C (Fig. 4.13). Interestingly, the use of cold C2 fragments caused
a supershift of the HrpY-probe C complex, suggesting the formation of
HrpY oligomeric complexes exposing multiple DNA-binding surfaces.
4.3.5 Binding of HrpY to 5’ hrpL DNA fragments
The binding of HrpY to the hrpL regulatory region was also studied by
electrophoretic mobility shift assays. Six DNA probes, spanning the
region from +221 to –148 (coordinates are relative to the transcriptional
start site of the σ 54-promoter), were synthesized by PCR (Fig. 4.14).
Probes were end-labeled with [γ32P]ATP, incubated with 20 µM HrpY in the presence of 1 µg of poly(dI:dC), and complexes at equilibrium were analyzed by native acrylamide gel electrophoresis (Fig. 4.15). HrpY retarded the mobility of probes L (+221 to –148) and M (-12 to –148), and to a lesser extent it bound to probes I (+221 to –94), H (+221 to –71), G
(+221 to –29). However, only a very small amount of probe F (+221 to –
11) was retarded. Titration experiments to determine the first approximation Kd were performed with the full-length hrpL promoter
(probe L). These results indicate that there is at least one strong operator site for HrpY, located between –12 and –29, and possibly a second weaker site(s) downstream of base -12. The specificity of the binding of HrpY to probe L was evaluated by competition experiments using specific and non- 208 specific competitor DNA (Fig. 4.16). As expected for specific binding, a
150-fold excess of poly(dI:dC) did not compete for the binding of HrpY to
probe L in the absence of poly(dI:dC), but a similar molar excess of cold probe L DNA did. The specificity of complex formation with the subfragments of probe L was not evaluated.
4.3.6 Effect of phosphorylation on in vitro HrpY binding activity
The effect of phosphorylation on the first approximation affinity of
HrpY for the target promoter sequences was analyzed by gel shift assays.
Probe C (Fig.C) was used to study the affinity for the hrpS promoter.
HrpY was phosphorylated either by BarA198 or by 50 mM carbamoyl phosphate, which is a small phosphodonor molecule often used as an alternative to acetyl phosphate. Increasing concentrations of phosphorylated (P) or unphosphorylated HrpY (from 5 nM to 80 µM) were incubated with a constant amount of probe C at nanomolar concentrations.
Protein-DNA complexes were separated on a 6% acrylamide gel and the bound fraction (ν) was measured by quantifying the amount of free probe
with a phosphor-imager scanner and comparing it to the no-protein
controls. The difference was used to calculate the bound fraction. Raw
data were transformed to linearize the binding isotherm and plotted in a
Scatchard graph (ν vs. ν/P) (Fig. 4.17A) and the apparent Kd was
calculated as the Y-axis intercept of the linear regression curve. When
HrpY was phosphorylated by BarA198, the Kd for binding to probe C was 209 decreased ca. 12-fold (from 20 nM to 1.7 nM). Similar changes in Kd
were obtained using carbamoyl phosphate. (Data not shown). These
results indicate that, at least in vitro, phosphorylation of HrpY can increase its affinity for target sites in the hrpS promoter region.
Similar experiments were done to measure the affinity of HrpY for the the hrpL promoter (probe L). However, it was necessary to use greater
concentrations of HrpY to obtain binding. The results in Fig. 4.17B show
that the Kd for binding to PhrpL is about three orders of magnitude higher
than that for binding to PhrpS. Interestingly, phosphorylation did not alter
the affinity of HrpY for its target sites in probe L.
4.3.7 Effect of phosphorylation of HrpY on its in vivo activity
The effects of conservative and structurally neutral substitutions at D57
in HrpY on hrpS expression were studied in P. stewartii. pRK415 plasmid
derivatives carrying Plac-hrpY[D57A] and Plac-hrpY[D57N] alleles were constructed as described in Chapter 2 (plasmids pMM74 and pMM118, respectively). A new hrpY mutant with a D57E substitution was engineered in vitro, following the SOE-PCR procedure described in
Chapter 2, and cloned into pRK415 to produce pMM99. The plasmids were mobilized into P. stewartii DM733, a strain carrying a chromosomal hrpS::lacZ fusion in a null hrpY background. The resulting strains were grown in IM5.5 and ß-galactosidase was measured (Fig. 4.18). Strain
+ DM733(pMM52), expressing Plac-hrpY , was used as positive control. The
210 HrpY[D57A] and HrpY[D57N] proteins were still able to activate hrpS, but its expression was only 33% and 25% that of the wild-type.
Interestingly, the HrpY[D57E] protein was completely unable to upregulate hrpS-lacZ.
4.3.8 Identification of the binding elements of HrpY by DNase I footprinting
The locations of the HrpY-binding sequences were identified by testing the ability of unphosphorylated HrpY to protect against DNase I cleavage of the of the hrpS promoter. Linear PCR with a 32P-labeled primer was used to amplify the partially digested complexes. Bottom strand-labeled fragments have not been tested yet. The experiments have not been repeated yet and the following results must be considered with caution.
Probe A (Fig. 4.10) was used for these experiments. Double autoradiography exposures were needed because the DNase I digestion ladders with the protein were underloaded (Fig. 4.19). A readable digestion ladder was obtained for sequences with coordinates +10 to –182.
Therefore most of the regions corresponding to probes B and C3 (Fig.
4.10) were not analyzed. Decreased DNase I sensitivity was shown for regions –50 to -71, -78 to –102, -125 to –165 and –174 to –182 (Fig. 4.19 black boxes). Hypersensitive sites were located around –50, -71, -104, -
106, -111, -123, and –138 (Fig. 4.19, solid arrows). These sites may indicate changes in the local bending angle of the hrpS promoter upon
211 binding of HrpY. The sequences protected by HrpY varied greatly.
Consequently chemical reagents with greater accessibility to the stacked
bases and/or the phosphate backbone of the DNA will be required to
identify a clear binding site. A tentative consensus site, MAATTYY,
shared by all the footprints is proposed (vertical arrowheads in Fig. 4.6).
This consensus exhibits dyad symmetry in the footprint located between –
78 to –102 compared to footprint at –165 to –148. When confirmed, these
results should support the specific binding observed with probe A and the
formation of three complexes between probe C2 and HrpY, but they may
contradict the absence of a mobility shift with probe C1. Potential
changes in the protection patterns upon binding by HrpY~P, which may be
critical in establishing a model for HrpY-dependent regulation of hrpS,
were not determined.
4.4 DISCUSSION
Data presented in this chapter provide the first detailed biochemical
analysis of the interaction of a hrp regulatory protein with its target
promoters. In particular, we provide evidence supporting the role of HrpY
as both an activator of hrpS and a repressor of hrpL. Possible models of
regulation are shown in Figures 4.20, 4.21 and 4.22.
As shown in Chapter 2 and Fig. 4.1, activation of hrpS is hrpY- dependent. Deletion of region –165 to –231 (corresponding to fragment B in Fig. 4.10) renders the hrpS-uidA fusion in pMM396 totally repressed
212 compared to the full-length fusion (e.g. pMM50 and pMM400). This
implies that DNA sites in fragment B, alone or in concert with other DNA
elements, are critically important for the activation of hrpS by HrpY. This is also consistent with the previous observation that hrpS loci lacking
fragment B are unable to complement null hrpS mutants, unless the gene
is driven by a strong vector promoter, such as Plac in pRF205 (Frederick
et al., 1993).
Electrophoretic mobility shift (EMS) assays were performed using
purified recombinant HrpY because crude cell extracts from P. stewartii
or E. coli showed nonspecific activity in negative control reactions (i.e.
null mutant strains or strains carrying the vector alone). This suggests
that bacterial proteins other than HrpY are able to bind to the probes (data
not shown). EMS assays showed weak, inconsistent binding of HrpY to
sites in fragment B (-165 to –231), no binding to fragment C1 (-90 to –
165), weak, erratic binding to C3 (+95 to –5) and stronger binding,
possibly at multiple sites, to fragment C2 (-5 to –90) (Fig. 4.10 and Fig.
4.12). Interestingly, probes A and C have much greater affinity for HrpY than the individual subfragments (approximate Kd <1 nM and 20 nM, vs.
micromolar ranges, respectively), as estimated visually from the bound
fractions in Fig. 4.11 and Fig. 4.12 or by Scatchard plot analysis (Fig.
4.17). This may be due to multiple discrete sites of interaction acting cooperatively. The fact that competition experiments using fragment C as
a probe and fragment C2 as the cold competitor created a supershift may
213 be interpreted as oligomerization of HrpY and interaction of some of the
subunits with non-contiguous DNA fragments. This also suggests that a
looping mechanism may be involved in hrpS activation in vivo (Fig.
4.21). Our preliminary DNase I footprinting experiments seem to indicate
that HrpY binds at multiple sites in the hrpS promoter region. These
experiments should be interpreted with caution at this point because we
have only examined the top strand protection and have not repeated the
experiment. Analyzing the top strand of probe A identified five footprints.
These sites span sequences corresponding to probes C2 and C1 and part of probe B (whose footprints were not well resolved by the experiments performed so far) (Fig. 4.19). Interestingly, one of the putative protected sites was split between probe C1 and probe C2, which may explain why the gel shift with probe C1 was negative (Fig. 4.12). Also, probe C2 contained two of the DNase I footprints and this finding is consistent with the multiple complexes seen in EMS gels. It is unclear at this point whether the weak binding to fragments B and C3 is specific or non- specific. The MAATYYY sites found in the regions protected by HrpY are quite degenerate, but for other FixJ-class response regulators, similar A+T rich sites have been reported, e.g. RAAAYY for UhpA (Dahl et al., 1997) and TACYNMT for NarL (Tyson et al., 1994). A more comprehensive footprinting analysis involving the use of chemical probes for major/minor groove and backbone interactions will be necessary to precisely define a clearer consensus binding site, especially in A+T rich
214 regions. Complex binding patterns at multiple sites were likewise
observed for E. coli NarL, a FixJ-class regulators, and OmpR, a “winged
helix” class regulator. In particular, NarL dimers (Maris et al., 2002) bind
four heptamers located in front of the fndG operon and to eight heptamers
in front of the narG operon in a cooperative and hierarchical manner (Li
et al., 1994). There may be different orientation and spacing among
heptamers in the various promoters regulated by NarL, but cooperative binding usually requires a central site with a 7-2-7 dyad symmetry
(Darwin et al., 1996). Another FixJ-class regulator, E. coli UhpA, binds as dimers to inverted repeats of the uhpT promoter, which were mapped by iron chelated hydroxy-radical footprinting (Olekhnovich and Kadner,
2002). On the other hand, asymmetrical OmpR dimers bind, in a cooperative fashion, multiple tandemly arranged 10-bp sites (Harrison-
McMonagle et al., 1999).
One possible working model consistent with the genetic and biochemical data invokes a repressor of hrpS acting on sequences located in fragment C1 (Figs. 4.20 and 4.21). This repressor would be required for maximal downregulation of the hrpS promoter. Its existence is hypothesized mainly to explain the observation that plasmid pMM396, containing the hypothetical operator site, has absolutely no promoter activity when compared to plasmids pMM393, which exhibits a low basal level, and pMM400, which is normally regulated by HrpY (Fig. 4.1). The absence of transcription termination demonstrated in Chapter 3 also calls
215 for a mechanism to downregulate read-through transcription of hrpY into hrpS under Hrp-repressing conditions. In this model, HrpY would then act as an antirepressor by binding to weak affinity sites in fragment B and to higher affinity sites in fragments C2 and C1, as shown by EMS assays
(Fig. 4.12) and the preliminary DNA footprintings (Fig. 4.19), respectively. The weak sites in fragment B have not yet been formally shown by footprinting but are suggested by the weak binding of HrpY to this probe. HrpY may therefore activate hrpS by forming a DNA loop involving both enhancer sites (at least three or four sites) and the site near the σ 70 promoter (Fig. 4.20). The presence of an IHF site
(AtgtAAccgttTTt) centered at –118 in fragment C1 further supports this looping model. IHF belongs to a class of DNA bending proteins that are able to impose strong curvatures (up to 180°) in A+T rich DNA sequences
(consensus WATCAAN4 TTR). Moreover, computer modeling suggests that
the hrpS promoter has as intrinsic curvature in the absence of any bound
factor due to the low G+C content and its estimated bending angle is 100°
(Fig. 4.4). In E. coli, IHF has also been shown to act as a repressor of
NarL-regulated genes (Browning et al., 2000). By analogy to the E. coli
Fis anti-activator (Browning et al., 2000), the hypothetical repressor
(perhaps IHF itself) may act upstream of the hrpS promoter and HrpY
would be the equivalent of the NarL antirepressor. (Wu et al., 1998).
Looping has been hypothesized for NarL dependent regulation of narK
(Kolesnikow et al., 1992), but probably it occurs via the formation of 216 higher order complexes of multiple transcription factors more than by
simple DNA bending (Dong et al., 1992). Alternatively, one might hypothesize that a second site, downstream of the σ 70 promoter, is
required for repression, which is typical of repressors in general (Wagner,
2000). This repression might likewise involve the formation of a DNA
loop (Fig. 4.21). Deletion of fragment B (-231 to –165) and C1 (–90 to –
165) would affect both the repressing mechanisms and the HrpY-
dependent activation of hrpS, as observed in Fig. 4.1 with plasmid
pMM393. The presence of repressor-binding sites in multiple upstream
positions has been described for the ompF gene in E. coli. In this system
both the –42 to –52 site and the –360 to –380 sites are important for
negative regulation of ompF, while sites in the –70 to –100 region are
important for activation of ompF by OmpR (Tsung, et al., 1989; Mizuno
et al., 1988; Ostrow et al., 1986). Both hypotheses are plausible, but other
explanations could be possible. Further research is needed to clarify the
mechanism used by HrpY. For example, in vitro transcription studies
and/or electron microscopy analysis of the hrpS promoter region using
purified HrpY and fractionated P. stewartii cell extracts may directly
demonstrate the existence of the repressor and the loops. Linker-scanning
or SOE-PCR mutagenesis could also be attempted in order to substitute or
delete the hypothesized operator site, ultimately confirming the DNA
binding data.
217 HrpY was also shown to repress transcription of hrpL (Fig. 4.22). The
basal expression of this gene was repressed about 7-fold by HrpY in an E.
coli reconstituted system (Fig. 4.2; pDM2785) and this repression did not require the UAS element recognized by HrpS (Fig. 4.2; pMM191).
Deletion analysis of the regulatory region of hrpL is not complete, but our
data to date suggest that an operator site downstream of base –12 is
required for the repression by HrpY (Fig. 4.2; pMM428). The
upregulation observed with plasmid pMM428 in the presence of hrpY is
not understood at this point. These data partially agree with the gel shift results, in that at least two sets of binding sites could be recognized: a high affinity site located between –29 and –12 and a potential weaker site located downstream of base –12. The specificity of the binding of HrpY to the latter site was not tested and the weakly shifted complex shown in lane two of Fig. 4.15 needs to be confirmed by further analysis. These two sites may act in concert and their affinity for HrpY may be decreased when they are located in separate probes. DNA footprinting of this region should provide more definite information on the location and specificity of the binding sites protected by HrpY and determine whether or not this is an overexpression artifact. What is notable is that the affinity of HrpY for the hrpL promoter is about 1000-fold lower than for the hrpS promoter
(Fig. 4.17). Assuming that binding constants measured in vitro are relevant to what happens in vivo, repression of hrpL would only occur
when HrpY reaches ca. 20,000 to 50,000 copies per cell, as compared to
218 the 20 to 50 copies needed to activate hrpS. From a biological point of view, hrpY may have acquired the additional function of fine-tuning the level of HrpL, a critical sigma factor coupling the transcription of all the type III secretion genes and their effectors. By indirectly activating hrpL via HrpS or directly repressing it, HrpY could quickly adapt the Hrp response to the environmental conditions sensed by HrpX. In P. stewartii, the HrpY regulator may also need to repress hrpL to act as a brake or the governor for the unique autoregulatory loop under non-inducing conditions. However, for this model to work, HrpY would also have to switch to repressing hrpS at the elevated concentrations needed for it to repress hrp. This mechanism of activation/repression would be similar to the regulation of ompF by OmpR (Rampersaud et al., 1994).
This study also explored the role of phosphorylation in modulating the activity of HrpY as a transcription factor. Protein phosphorylation gels
(Fig. 4.9) showed that the D57 residue is probably the sole phosphorylation site. More sensitive techniques, such as phospho-aspartyl derivatization followed by MS/MS spectrometry (Sanders et al., 1992), were not attempted. For other response regulators, phosphorylation affects their activity as transcription factors in various and often overlapping ways; it may change their quaternary structure, by promoting dimerization or oligomerization (Han and Winans, 1994; Li et al., 1994; Fiedler and
Weiss, 1995; Mettke et al., 1995), their DNA binding affinity to single or multiple cooperatively-acting sites (Galinier et al., 1994; Rampersaud et
219 al., 1994; Huang and Igo, 1996; Dahl et al., 1997; Huang et al., 1997;
Bergstrom et al., 1998). Our results showed that phosphorylation of HrpY increases its affinity for DNA sites in the hrpS promoter (Fig. 4.17 A), but not in the hrpL promoter. More precise assessments of the binding affinities using filter-binding assays or electron-plasmon resonance were not pursued. Consequently, we consider the Kds calculated from Scatchard plots as “first approximation” constants. The importance of D57 in transcription activation was also tested in vivo using unphosphorylatable hrpY[D57] alleles expressed from multicopy plasmids. The reduction in hrpS activation due to the hrpY[D57N] and hrpY[D57A] mutations, as compared to wild type hrpY, are probably due to the effect of these mutations on the Kd , or, in other words, to the inability of HrpY[D57N] to oligomerize at the hrpS promoter to the same extent as HrpY~P. However, it could alternatively reflect the inability of the unphosphorylated form to fully activate the RNA polymerase holoenzyme. Genetic screening for positive control mutations in hrpY, i.e. those affecting activation but not
DNA binding, together with screenings for suppressing mutations in components of the RNA polymerase holoenzyme may be helpful in separating these two effects more clearly (Aiba et al., 1994). In this regard, the hrpY[D57E] mutant may be an interesting mutant to study in more depth with biochemical approaches because molecular modeling predicts it should produce a correctly folded protein as competent for
DNA binding as HrpY[D57N], but yet, it is unable to activate hrpS, even
220 when overexpressed. In other systems, similar substitutions at the
phosphrylation site render the response regulator constitutively active
(Klose et al., 1993).
Salmonella enterica sensor kinase truncated mutant BarA198 (Teplitski
et al., 2003) was used to phosphorylate HrpY in vitro, both in protein labeling experiments and in gel shift assays. BarA orthologs are found in many γ-proteobacteria, including the genera Pseudomonas, Pantoea,
Erwinia and Pectobacterium. Chatterjee and coworkers (Cui et al., 2001) used low stringency Southern blot hybridization to show that P. stewartii also has a barA/gacS-like gene. Usually there is a certain degree of specificity between response regulators and their cognate kinases in order to achieve proper regulation (Hoch and Varughese, 2001). In particular, E. coli BarA is quite specific for its cognate UvrY regulator (Pernestig et al., 2001). Whether the phosphorylation of HrpY by BarA198 described in this study is an artifact of the elevated protein concentration used in vitro or it has physiological relevance in P. stewartii cells is not known. It is worth noting that BarA/SirA and GacS/GacA have important roles in virulence regulation of salmonellae, erwinias and pseudomonads.
Small phosphodonors can phosphorylate response regulators in vitro and in vivo (Lukat et al., 1992). In this study, carbamoyl phosphate was shown to modify the affinity of HrpY for hrpS promoters and this was interpreted as an indication of its ability to phosphorylate HrpY. Acetyl phosphate did not show the same effect (not shown). Formal
221 demonstration of this reaction using radiolabeled carbamoyl phosphate was not attempted because we did not have the means to synthesize it.
Future studies on the alteration of HrpY footprints at the hrpS and hrpL promoters might use this convenient and cheap small phosphodonor instead of recombinant kinases.
In conclusion, the dual role of HrpY as an activator of hrpS and a repressor of hrpL was supported in this study by independent approaches.
However, the biological significance of and the inherent logic behind its role as a repressor remains somewhat puzzling and open to different interpretations. Our preliminary data on the interaction of HrpY with specific sites in the hrpS promoter represents an important step toward the future in vitro analysis of other global and specific transcription factors in controlling this key step of the hrp regulatory cascade. These factors include: a) the DNA-bending protein IHF; b) the quorum sensing response regulator EsaR; c) the DNA-binding protein H-NS, which we have shown binds to probe C (Merighi and Coplin, unpublished data); and d) other novel regulators, such as a NikR-homolog, a member of the ribbon-helix- helix family of transcription factors recently implicated in the negative regulation of hrp genes (Ham and Coplin, unpublished data).
222
4.5 SUMMARY
Pantoea stewartii subsp. stewartii is a bacterial pathogen of corn,
whose pathogenicity depends on translocation of effector proteins into host cells by the Hrp/type III secretion system. Using genetic approaches, we delineated a regulatory cascade that controls the hrp/hrc secretion and wts effector genes in response to environmental signals and various global regulators. The cytoplasmic HrpX/HrpY PAS kinase/response regulator pair senses an unknown internal signal resulting in the phosphorylation of
HrpY. The most likely phosphorylation site in HrpY is a conserved D57 residue that is required for virulence. HrpY activates hrpS transcription
and HrpS, in turn, enhances transcription of hrpL. At the end of the
cascade, HrpL is an ECF alternative sigma factor that activates the genes
in the Hrp regulon.
In this study, we characterized the regulatory elements in the hrpS and
hrpL 5’ regions using primer extension analysis, electrophoretic mobility
shift assays and a series of overlapping subclones creating uidA reporter
gene fusions. Progressive deletions from the 5’ end of the hrpS promoter
region first abolished transcription and then restored it, but it was no
longer HrpY-dependent. This finding is consistent with the deletion of an
operator for an unknown transcriptional repressor and a model in which
HrpY acts as an antirepressor. A similar analysis of the hrpL promoter has
yet to be completed, but our experiments to date suggest the presence of
223 an operator required for HrpY-dependent repression downstream of the
54 σ promoter. Purified, unphosphorylated His6 -HrpY bound specifically to portions of the 5' regulatory regions of hrpS and hrpL in gel retardation assays. In vitro phosphorylation of HrpY increased its binding affinity for the hrpS promoter, but not for the hrpL promoter. Preliminary electrophoretic mobility shift assays and DNase I footprinting experiments suggest that HrpY may bind to multiple sites within each promoter region.
224 Strain or plasmid Relevant phenotypes and genotypes Source or reference
Escherichia coli
DH10B F- mcrA ∆(mrr-hsdRMS-mcrBC) φ80dlacZ∆M15 ∆lacX74 endA1 recA1 deoR
∆(ara,leu)7697 araD139 galU galK nupG rpsL λ- Invitrogen
- - - r HB101 F thi-1 hsd20 (r Bm B) sup E44 recA13 ara-14 leuB6 proA2 lacY1 rpsL20 (Sm )
xyl-5 mtl-1 Boyer and Roulland-Dussoix (1969)
Sφ200Rif metB strA purB ∆(agg-uidA-man) Rifr Wei et al. (2000)b
BL21(DE3)/pLysS F- ompT hsdSB (rB- mB-) gal dcm (DE3) caarying pLysS Novagen
P. stewarti subsp. stewartiii
DC283 Wild type, Wts+, HR+ Nalr Coplin et al. (1986)
DM064 DC283 hrpY1296::Tn5 (Kanr) Frederick et al. (2001)
MM254 DC283 hrpY(D57N) Merighi et al. (2003)
Plasmids pBluescript
KS and SK (+) ColE1 αlacZ (Apr) Stratagene pRK415 IncP αlacZ (Tcr) Keen et al. (1988)
pPL6GUSC pLAFR6 derivative carrying a promoterless uidA gene (Tcr) Knoop et al. (1991)
pMM221 pET15b with a 0.7 kb NdeI hrpY PCR fragment This study pMM222 pET15b with a 0.7 kb NdeI hrpY D57N PCR fragment This study
pMM46 pBKS(+) with a 0.7 kb HindIII hrpY fragment made by PCR, transcribed from Plac Chapter 2 pMM92 pBKS(+) with a 0.7 kb HindIII hrpY(D57N) fragment made by PCR-SOE This study pMM50 pPL6GUSC with a +994 to –317 hrpS PCR fragment fused to uidA Chapter 2 pMM400 pPL6GUSC with a +629 to –231 hrpS PCR fragment fused to uidA This study pMM396 pPL6GUSC with a +629 to –165 hrpS PCR fragment fused to uidA This study pMM393 pPL6GUSC with a +629 to –90 hrpS PCR fragment fused to uidA This study pMM403 pPL6GUSC with a +629 to –51 hrpS PCR fragment fused to uidA This study pMM392 pPL6GUSC with a +629 to –5 hrpS PCR fragment fused to uidA This study pDM2785 pPL6GUSC with a +220 to –148 hrpL PCR fragment fused to uidA Chapter 2 pMM191 pPL6GUSC with a +220 to –95 hrpL PCR fragment fused to uidA Chapter 3 pMM428 pPL6GUSC with a -12 to –148 hrpL PCR fragment fused to uidA This study
Table 4.1. Bacterial strains and plasmids.
225
Sequence coordinates Primer Sequence (5’Æ 3’) (5’Æ3’)a Source
ET-YF-NdeI GGGAATTCCATATGGATAACACGATTCGAATA 2866-2887 This study
ET-YR-NdeI GGGAATTCCATATGCAATCAGTCAATTAAGAGCAATCCTAAGAGA 3501-3525 This study
JFBG aka HrpLP1 CGCGGATCCGAGCTCTTTCCCAGGCAACAGAGTTAA 481-501 Merighi et al., 2003
L578F CGCGGATCCAATGTAAATCGTTAAATTTC 578-597 This study
L597R CGGGGATCCTGAAATTTAACGATTTAC 581-597 This study
L620F CGCGGATCCGCTGGCACAGACTTTGCTAT 620-639 This study
L637R CGGGGATCCAGCAAAGTCTGTGCCAGCGA 618-637 This study
L638F CGGGGATCCATCGCCATTAAGCCAAACGG 638-657 This study
LRBG CGCGGATCCTCTAGAATGACTTCCAGCCAGGTCAT 869-850 Merighi et al., 2003
PEL TATGTGGGGATGGAGGGTTAACGTG 724-748 This study
SF3509B CGCGGATCCGATTGCTCTTAATTTACAAAT 3509-3529 This study
SF3575B CGCGGATCCAAAAAAGCTTTTTAAACAGC 3575-3594 This study
SF3650B CGCGGATCCGAAATCCTTACAATCCTC 3650-3667 This study
SF3689B CGCGGATCCCAGCATATGACGATGGTTT 3689-3707 This study
SF3735B CGCGGATCCCCCCTGTAAGCACTTCGTAA 3735-3754 This study
SFBG CGCGGATCCATAAAGCCCGCCTGATGGAA 3423-3442 Merighi et al., 2003
SR3575 GCATAAGAAATACCATGTC 3717-3735 This study
SR3650 CCTGACTAACCCTGTGTGAAATG 3628-3650 This study
SR3735 GCATAAGAAATACCATGTCA 3817-3835 This study
SR4368B CGGGATCCGCTCACTCACTCACATTATTTTCAATATTCAT 4355-4369 This study
YD57E TCTTTTACTTCTAGAAATGTGTATGCCTGG 3022-3051 This study
YFH CCGAAGCTTTTCGATAACAATATGGATAACAC 2855-2877 Merighi et al., 2003
YR2 GTTTCAGGATCGTTATCTTTCGTGATG 3204-3178 Merighi et al., 2003
YRH CCGAAGCTTATTAAGAGCAATCCTAAGAGA 3521-3501Merighi et al., 2003
Table 4.2. Oligonucleotide primers.
226
RBS +1 σ70 +611 GUS activity HindIII hrpS IS-like hrpY
WT HrpY[D57N] +994 -317 214±26 16 ±4 gusA pMM50 +629 -231 317 ±9 98 ±2 gusA pMM400 +629 -165 gusA pMM396 1±0 1±0.5 +629 -90 70±5 68±2 gusA pMM393
+629 -51 99±24 79±6 gusA pMM403
+629 -5 48±7 39±1 gusA pMM392
Figure 4.1. Expression of plasmid-borne hrpS gene fusions in P. stewartii.
Expression of GUS reporter plasmids shown below the map (the coordinates of the
sequence ends are shown on the map relative to the transcription start site) was
analyzed in strains DC283 (wild-type) and MM254 (DC283 hrpY[D57N]) after growth
in IM5.5 for 16 h. Data are means from two experiments with two replicates each.
-1 –1 70 GUS activities are reported as pmol MU min OD600 ± standard deviation. The σ promoter is indicated by a yellow vertical arrow. RBS = ribosomal binding site.
227
+1 σ54 σ70 σL IHF UAS GUS activity hrpL hrpJ Control +hrpY +220 -148
76 ±9 5 ±1 gusA pDM2785 +220 -95 156±10 15±8 gusA pMM191 -12 -148 4±3 53±13 gusA pMM428
Figure 4.2. Expression of plasmid-borne hrpL gene fusions in E. coli SΦ200.
Expression of GUS reporter plasmids shown below the map (the coordinates of the
sequence ends are shown on the map relative to the transcription start site) was
analyzed in the E. coli strain SΦ200, with and without the regulatory plasmid pMM46
+ P lac-hrpY , after growth in LB broth for 16 h. Data are means from two experiments
-1 –1 with two replicates each. GUS activities are reported as pmol MU min OD600 ± standard deviation. The positions of σ 70, σ 54, and σ L promoters in the hrpJ-hrpL intergenic region are shown by yellow arrows. IHF = integration host factor; UAS = upstream activating sequence.
228
Figure 4.3. Nucleotide sequence alignment of the hrpY-hrpS intergenic regions
from several erwinias and salient features of this region in P. stewartii.
The DNA sequence of the P. stewartii (Pns) hrpS regulatory region minus the 483 bp
IS-like element was aligned with the corresponding regions from P. agglomerans pv.
gypsophylae (Pag), Erwinia amylovora (Ea) and Pectobacterium chrysanthemi (Pch)
using ClustalX. Primers for probes and plasmid fusions are indicated with a name
followed by a small arrow; the coordinates in the primer names refer to P. stewartii
GenBank accession AF282857. Regions protected in footprinting experiments
described in Fig. 4.6 are shown here in bold italics, and the thick arrows indicate the
putative HrpY consensus binding site (MAAYYY). Peaks present in the intrinsic
curvature profiles shown in Fig. 4.4 are marked by asterixes. RBS = ribosomal
binding site; UTR = untranslated region; CUR = Conserved upstream region; VUR =
variable upstream region.
229 SF3509B →
CUR
Pns GATTGCTCTTA-ATTTACAAATTAATGCTACCCACCGTTATTACTTTATCATTTCAACAG Pag GATTGCTCTTATATTTTTTTATTAATGCTACCCATCGTTATTATTTAATTATTTCAATGC Ea GAATGCTCTTATATTTGTCT-CTCGCCCTTCCCTCCTTGAGCAGATATTTATTATCACGC Pch ------CGGTA-ACTTGTACGC---CGCTCCTCTCCGTTATGCGTCATCGCCTGTCGTGA
HindIII VUR
SF3575B → ****** ******* IHF site
Pns GTTACAAAAAAAAGCT-TTTTAAACAGCACGTTATGCCTAAAAATTTTATGTAACCGTTT Pag TTTA-ATAAAAACCCT-TTATTGACAACCTGTTATGTTCATCG-CGATGTGTAAACGCTT Ea CAGATTTAAAAACACCATTAAAAACAATTGGATAAAATGGTTG-TGGAGTGTAACCGCTT Pch CGGAGAGGAGATCCTCGTTTCCCCCTATCCGGAGAGCC------TGCGTAGCGGCG-
Direct Repeats
←SR3650B SF3650 →
Pns TCATTTCACACAGGGTTAGTCAGGAAATCCTTACAATC--CTCCATCAATTCCTTACCCG Pag CCACTCTAATGATGGTGGGTAGGAATAGCCTTACCCTTATCTCAGTGAATTCCTTACTCG Ea CCATTTTATCCCCCGAATGTAGGGTAATCCCTACATTGCCCCCTGAAGATTCCTTACTTA Pch ------ATTTCCGCGGGGTAAGATTATCCCTACTTTTTGCTCTG--AAATCCCTACCTC
σ70 promoter
SF3689B → -35 box -10 box ← SR3735 SF3735B→
Pns ATAT-CAGCA-TATGACGATGGTTTATCT-GTGCTGACATGG-TATTTCTTA-TGCCCCT Pag TGGTTCAGCA-TAAGACGATGGTTAATCT-GAGCTGGCATGA-TATTTTTTA-ATTCATT Ea ATGTTCAGCA-TAAGACGATGGTTTCTGG-ACGTTAGTATGGGTATTTCTTAGTGGCAGC Pch AACTTCAGCGGTTGGCTGATGGTTTGTTTTGCGCCAG-GCAG-TAATGTTTACGCATACC
mRNA start +1 5' UTR
Pns GTAAGCACTTCGTAAACATAAGAATAGTGATTGTAATTAAATGTAATTCAGCTATTGGCT Pag GTAAGCGAAGCGTAATCACGTGTGCAGTCAGTGTAATTAATTGTAAGTGATGAGGTGGCA Ea GTAAACTCAGAGTAAATATTACAGGCATAATTGTAAATAGTTGTGTCTTGTATAACGGCT Pch ATCAATACGTCGTTAAAACGATCAGATTTATCGCAGTTGATCGTTAAGAATACATCAGGG
Site of 483-bp IS-like insertion in Pns ←SR3845
Pns CTAAGTATGTTGTT--AACTGCGATGATGTCAGTTGAAA----GGATAAGTG------Pag ATTAGATTGCTGTG--TACCGCAATGCCGTTAAATAAAACA--GGATAGGAA------Ea GTAACATGACTGTGC-AAGTGAAGCAATGTTAAAAAAAATAATGGATAGGGC------Pch TGATGATGATGGTTGTCAGATCGACGCTGTCGCCGACATGTCGGCACGGCATGGAAATGA ** * ** * * *
RBS Start codon Pns ------AGGGGCAACTTTTTATG Pag ------AGGGGTAACTTCTTATG Ea ------AGGGGTAATTTTTTATG Pch TCCGACAGTCTGATAATGGAATACAAAAAAAGTACAGGAGCAGTC----ATG
230 -231 -154 +1 +95
Figure 4.4. Computer modeling of the intrinsic curvature of the hrpS promoter
region.
The DNA sequence of the hrpS promoter region corresponding to fragment A in Fig.
4.10 was analyzed with the Bend.It algorithm (www.icgeb.trieste.it) to estimate the
intrinsic curvature of the DNA molecule due to its base composition. The graph shows
the predicted curvature in degrees per helical turn and the G+C content in % using a
nucleotide window of 31. Coordinates in bold are relative to the transcription start.
The double-headed arrow indicates the region integrated to calculate the total
bending.
231
Figure 4.5. Primer extension analysis and sequence analysis of the hrpL promoter.
(A) Total RNA from DC283 cells grown in IM5.5 or LB broth was annealed with the
32P-end-labeled primer PEL and cDNA was reverse transcribed in vitro. Lanes A, T,
G, and C show the dideoxy sequencing ladders for the hrpL gene generated with the same primer. The transcriptional start sites are indicated by solid arrows. The transcription starts and the σ 54 promoter region are highlighted. (B) Sequence analysis
of the hrpJ-hrpL intergenic region. Coordinates are relative to Genbank accession
AF282857. Promoter boxes are in bold, underlined characters. RBS = ribosomal
binding site.
232
A
LB IM5.5 A C G T
…AAACCG AA TT A CCGCTATCGTTTCAGACACGGTCGCTAAAATAAAACAA TTTGT AAACTTTAAA +1 σ54 -58 (+1’) B
421 CTCATACAGGGAAGCTGATGCAGATATGTGGCGTGTTTTATGCTCCGGTTCCCAATAAAT 480 R M ← hrpJ UAS
481 TTTCCCAGGCAACAGAGTTAATTACAGAAGATTGCAATGAGTTGCAGATTTAGCTGGAGC 540
sigma 70 box -58
-35 -10 +1’ 541 TTACGGCAAAAAGTTGCGAAAAAATGAAGGGAAATAAAATGTAAATCGTTAAATTTCAAA 600
IHF site sigma 54 box
-24 -12 +1 601 TGTTTAACAAAATAAAATCGCTGGCACAGACTTTGCTATCGCCATTAAGCCAAACGGCAA 660
RBS__ 661 AAAATGGATTAAGATACGGAGCGTATTATGTCAGAAATGAACCTGCACCCTACTGAGTCA 720 M S E M N L H P T E S hrpL →
233
Figure 4.6. Overexpression and purification of HrpY.
Plasmid pMM221 was expressed in E. coli BL21(DE3)/pLysS upon IPTG induction.
Proteins were purified by native IMAC on Ni2+-IDT columns. Various protein
fractions were separated by SDS-PAGE and stained with Coomassie Blue. (A): lane 1,
total cell proteins of uninduced cells; lane 2, total cell proteins of IPTG-induced
cells; lane 3, flow-through from the Ni2+-IDT columns loaded with a French press
lysate; lanes 4 and 5, fractions eluted by using washing buffers with 20 to 60 mM
imidazole; lane 6, 250 mM imidazole fraction; lane 7, 500 mM imidazole fraction.
(B): Step gradient purification fractions with elution buffers containing imidazole at
150 mM (lanes 1 to 3), 250 mM (lanes 4 to 6) and 250 mM (lanes 7 to 9). Fractions 1-
2 and 3-9 were separately pooled, desalted by G25 chromatography and purified by
ion exchange chromatography. (C), fractions purified by anion exchange
chromatography on Q-sepharose columns. Fractions from lanes 5 to 8 were pooled and
desalted/concentrated by ultrafiltration. Proteins purified by ion-exchange
chromatography were used for all the phosphorylation experiments and for most of the
DNA binding assays.
234 123 45 6 7 A 61.3 kDa 49 36.4 24.7 19.2 13.1
9.3
1 2 3 4 5 6 7 8 B 61.3 kDa 49 36.4 24.7 19.2
13.1
9.3
1 2 3 4 5 6 7 8 9 10 11 12 13 14 kDa 61.3 C 49 36.4 24.7 19.2
13.1
235
A B
C D
Figure 4.7. Mass spectrometry analysis of recombinant HrpY and HrpY[D57N].
Protein molecular weight measurements on 300 pmol of desalted HrpY and
HrpY[D57N] were performed by electrospray ionization mass spectrometry using a
Micromass Q-TOFTM II (Micromass, Wythenshawe, UK) mass spectrometer at the OSU
CCIC-MS Facility. m/z spectral data for HrpY (A) and HrpY[D57N] (C) were deconvoluted in the 22 to 30 kDa region using MaxEnt software (Micromass) to
produce distributions of molecular weights for HrpY (B) and HrpY[D57N] (D).
236
Figure 4.8. Homology model of the crystal structure of P. stewartii HrpY
A structural homology model of the HrpY protein was constructed with the Swiss-
Model server using the X-ray diffraction and NMR solution structures of E. coli NarL and Bacillus subtilis Spo0F. Modeling was successful from residue 3 to 206 and
AcelrysView software was used to render the structure. The modular structure of the
protein is shown in panel A. Details of the catalytic site with the conserved
aspartates, lysine and threonine are shown in panel B. Panel C shows the LuxR-like
helix-turn-helix (HTH) motif in the output domain of HrpY. The complete structural
coordinates are reported in the Appendix.
237
Receiver domain Linker Catalytic triad
Output domain
A
D57 D11
D12 K105
T85
DNA binding Helix-turn-helix motif
B Catalytic site residues C
238
HrpY - + - + - - + - + - HrpY[D57N] + - - - + + - - - + BarA - - + + + - - + + +
BarA198~P
26 kDa HrpY~P
Coomassie Autoradiogram
32 32 Figure 4.9. Phospho-transfer of P i from BarA198~ P to HrpY.
Reactions contained 25 µM HrpY, 75 µM Salmonella enterica BarA198, 40 µM
[γ 32P]ATP and 0.1 mM cold ATP in a total volume of 15 µl. Mixtures were incubated
at 25°C for 90 min. Reactions were terminated with 5 µl of 4X stop buffer, separated
by SDS-PAGE (T=12%), and visualized by Coomassie Brilliant Blue staining.
Radiolabeled proteins were visualized by Phosphor-Imager scanning before gel
staining. Negative controls consisted of HrpY and HrpY[D57N] incubated in presence
of [γ 32P]ATP.
239
70 +1 σ
hrpS IS-like hrpY
+95 -231 A 327 bp
-165 -231 B 76 bp +95 -165 C 261 bp -5 -90 C2 86 bp +95 -5 C3 101 bp -90 -165 C1 76 bp
Figure 4.10. Map of the hrpS regulatory region and the promoter fragments used as probes in gel shift experiments
The hrpS promoter region was amplified by PCR from plasmid pMM58 to produce fragments A, B, C, C1, C2 and C3. Coordinates are relative to the transcription start site. Fragments were used in electrophoretic mobility shift experiments as either radiolabeled probes or cold specific competitors.
240
hrpS probe (25 fmol) : A
HrpY (nM): 0 10 40 160 625 625
Competitor (A fragment): - - - - - +
261 bp
Figure 4.11. Electrophoretic mobility shift analysis of the binding of HrpY to the full length hrpS promoter.
Binding conditions are described in Materials and Methods. Increasing concentrations of HrpY were incubated with 25 fmol of 32P-labeled fragment A DNA (Fig. 4.10) for
35-40 min at room temperature. Aliquots were loaded onto a 5% acrylamide gel in
1XTBE buffer and electrophoresed for 2.5 h at room temperature. The positions of the
free probes and complexes were determined by autoradiography of the dried gel at –
80°C. The molar excess of the specific cold competitor in the last lane over the 32P- probe was 100-fold. The size of the free probe in bp is shown.
241
hrpS probe B C1 C2 C3 C (10-20 fmol) : HrpY(4.7 µM): - + + - + + - + + - + + - + + + Competitor (C fragment): - - +200X - - +62X - - +114X - - +46X - - +5X +50X
261 bp
103 bp 86 bp 76 bp
Figure 4.12. Electrophoretic mobility shift analysis of the binding of HrpY to portions of the hrpS promoter.
Binding conditions are described in Materials and Methods. Assay mixtures contained
4.7 µM HrpY and 10-20 fmol of 32P-labeled probe DNA (10,000 cpm). Fragments B,
C, C1, C2 and C3 (whose coordinates are reported in Fig. 4.10) were used as probes.
Mixtures were incubated for 35-40 min at room temperature. Cold competitor DNA
(probe C) was added before HrpY at the concentrations indicated by superscripts
(expressed as molar fold over the 32P-probe). Aliquots were electrophoresed in a 5%
acrylamide gel in 1X TBE buffer for 2.5 h at room temperature. The positions of the
free probes and complexes were determined by exposure of the dried gel to an X-ray
film at –80°C. Sizes of the free probes in bp are shown.
242
hrpS probe: C
HrpY (12.5 µM): - + + + + + + + + + + + + + + + +
Competitor -- BC1C2C3C (25,100,250X):
Figure 4.13. Competition analysis of the binding of HrpY to hrpS fragment C
Binding conditions are described in Materials and Methods. HrpY at 12.5 µM was
incubated with 25 fmol of 32P-labeled DNA fragment C (coordinates reported in Fig.
4.10) with and without competing unlabeled DNA fragments (B, C, C1, C2, C3; see
Fig. 4.10 for the coordinates) at 20, 100 and 250 molar excess for 35-40 min at room temperature. Aliquots were electrophoresed in a 5% acrylamide gel in 1XTBE buffer for 2.5 h at room temperature. The positions of the free probes and complexes were determined by exposure of the dried gel to an X-ray film at –80°C. Sizes of the free probes in bp are shown
243
54 70 L + σ σ σ UAS hrpL hrpJ
+221 -148 L389bp
+221 -94 I 315 bp
+221 -71 H 292 bp
+221 -29 G 250 bp +221 -11 F 232 bp -12 -148 M 156 bp
Figure 4.14. Map of the hrpL regulatory region and of the promoter fragments used as probes in gel shift experiments.
The hrpS promoter region was amplified by PCR from plasmid pMM58 to produce fragment L with the complete promoter region fragments I, H, G and F with 5’ deletions, and fragment M with a 3’ deletion. These were used in electrophoretic mobility shift experiments. Sizes are in bp and coordinates are relative to the transcription start site. UAS = upstream activating sequence.
244
hrpL probe (10-25 fmol) : F G H I L M
HrpY (20 µM): - + - + - + - + - + - +
389 bp 292 bp 315 bp 250 bp 232 bp
156 bp
Figure 4.15. Electrophoretic mobility shift analysis of the binding of HrpY to portions of the hrpL promoter.
Binding conditions are described in Materials and Methods. HrpY at 20 µM was
incubated with 10 to 25 fmol of 32P-labeled DNA (5,000 cpm) corresponding to fragments F, G, H, I, M and L (whose coordinates are reported in Fig. 4.14) for 35-40
min at room temperature. Aliquots were electrophoresed in a 5% acrylamide gel in 1X
TBE buffer for 2.5 h at room temperature. The positions of the free probes and
complexes were determined by exposure of the dried gel to an X-ray film at –80°C.
Sizes of the free probes in bp are shown.
245
hrpL probe: L
Aspecific Specific Competitor(fold):
-- - - -1x 7X 25X 150X 1X 7X 30X 155X
HrpY(µM): 0 1 10 20 50 5050 50 50 50 50 50
389 bp
Figure 4.16. Electrophoretic mobility shift analysis of the binding of HrpY to the full length hrpL promoter fragment.
Binding conditions are described in Materials and Methods. Increasing concentrations of HrpY were incubated with 5 fmol of 32P-labeled DNA corresponding to fragment A
(6000 cpm) in Fig. 4.10 for 35-40 min at room temperature. Specific or aspecific competitor DNAs consisted of unlabeled DNA fragment L and poly(dI:dC), respectively. Aliquots were electrophoresed in a 5% acrylamide gel in 1X TBE buffer for 2.5 h at room temperature. The positions of the free probes and complexes were determined by exposure of the dried gel to an X-ray film at –80°C. Sizes of the free probes in bp are shown.
246
hrpS probe C hrpL probe 1E+08 8000 A B 9E+07 7000 8E+07 HrpY Kd=20 µM HrpY Kd=20 nM 6000 HrpY~P Kd=21 µM 7E+07 HrpY~P Kd=1.7 nM
6E+07 5000
5E+07 4000 ν/P HrpY~P ν/P HrpY~P 4E+07 3000 3E+07 2000 2E+07 HrpY 1000 HrpY 1E+07
0 0 0 0.5 1 0 0.5 1 ν ν HrpY HrpY~P HrpY HrpY~P
Figure 4.17. Effect of phosphorylation on HrpY binding to the hrpS and hrpL promoters
Increasing concentrations of HrpY or HrpY~P (ranging from 5 nM to 80 µM with concentrations increased 4-fold each step; first lane in each series is the “no protein” control) were incubated with an average of 40 fmol of 32P-labeled probe DNA (10,000
cpm) for 35-40 min at room temperature in binding buffer (see Materials and
Methods). Aliquots were loaded onto a 5% acrylamide gel in 1X TBE buffer and
electrophoresed for 2.5 h at room temperature. The positions of the free probes and
complexes and their intensities were determined by exposing the dried gel to a
phosphor storage screen (lower panels). (A): Binding of HrpY to hrpS fragment C
(similar results were obtained for fragment A). (B): Binding of HrpY to hrpL fragment L (coordinates are reported in Fig. 4.10 and Fig. 4.14).
247
DM733/pMM118 D57N
DM733/pMM99 D57E
DM733/pMM74 D57A
DM733/pMM52 D57
DM733
0 100 200 300 400 500 600 β-galactosidase activity (Miller Units)
Fig. 4.18. β-galactosidase activity of a chromosomal hrpS-lacZ fusion in a P. stewartii hrpY null mutant carrying various plasmid-borne Plac-hrpY regulatory
genes
Cells were grown in IM pH 5.5 broth to late log phase and β-galactosidase specific
-1 -1 -1 activity was measured (1 unit = 1 pmol MU min OD600 ml of culture at 25°C).
+ Regulatory gene clones: pMM52 Plac-hrpY , pMM74 Plac-hrpY[D57A], pMM99 Plac-
hrpY[D57E], and pMM118 Plac-hrpY[D57N]. Strain DM733 is DC283 hrpY::Tn5
hrpS::Tn3HoHoI. Vector control not run in this particular experiment. Previous data
suggested no effect of pRK415 on P. stewartii hrp genes and no effect of recombinant
plasmids on DC283 β-galactosidase background activity (not shown).
248
Fig. 4.19. DNase I footprinting assay of HrpY complexes at the hrpS promoter.
Binding reactions used HrpY at the concentrations shown above the lanes and 35 fmol
of a DNA fragment spanning probe A. The equilibrated complexes were partially
digested with DNase I and the digests were amplified by linear PCR with 32P-labeled primer SF3509. The reactions were electrophoresed in a 6% acrylamide/8M urea sequencing gel alongside sequencing ladders for A and G. Solid arrows indicate hypersensitive sites, solid black boxes represent protected sequences. Lane “0” was loaded with 4 times the normal amount of DNA in order to visualize the ladder. The control ladders were visualized by autoradiography for 24 h and the footprint ladders required several days of exposure at -80°C. The coordinates are relative to the +1 transcription start. The bars on the right represent portions of the probe A.
249
HrpY A G0 1.5 15 µM
C3
+1 -10 box -35 box
-50 -50 C2 -71 -80 -78
-100
-102
-118 IHF -120 -125 C1 -132
-148
-165 -170
-174 B (not shown in -182 full)
250
A
R =IHF?=H-NS? RNApol IS-like C3 σ70 C2 C1 B +1 IHF
B
Hierarchical cooperative binding?
RNApol Y Y Y Y Y IS-like C3 σ70 C2 C1 B +1 IHF ?
Figure 4.20. Model for hrpS activation.
(A) Repression may involve a factor ( R ) binding at the C1 fragment. An IHF site is contained in this region. Whether IHF is a repressor of this system, as shown in other
E. coli promoters controlled by cooperative binding of FixJ-like regulators, or a helper for HrpY is not known. H-NS was also shown to bind to C fragments in preliminary experiments (not shown) (B) Cooperative, perhaps hierarchical, binding of HrpY (Y) at multiple sites activates the hrpS promoter. Given that binding to C1 and B is weak or not visible in EMS assays, but observable in the preliminary footprinting experiments, initial binding to C2 sites may be a prerequisite. This is consistent with the stronger binding of HrpY to the A probe compared to binding to its subfragments.
251
Activation loop
IS-like C3 C2
HrpY HrpY C1 B
Repression loop C3
IS-like R?
C2 R?
B C1
Figure 4.21. Model for hrpS activation.
The activation/repression model shown in Fig. 4.20 may involve looping of the hrpS promoter region, supported by the presence of an IHF site and by the intrinsic bending of the DNA.
252
A Activation loop
σ54 σ70
hrpL HrpS IHF
UAS
B Repression loop
hrpL
? -12 HrpY 54 σ
-29 σ70
Figure 4.22. Models for hrpL regulation.
(A) Activation of the σ54 promoter of hrpL may involve binding of HrpS to the UAS
element (probably as simple dimers because of the presence of a single dyad in the
site), DNA looping, perhaps helped by IHF, and isomerization of closed complex. (B)
DNA binding and repression by HrpY requires sites downstream and upstream of the –
12 base. This implies that a looping mechanism may be involved as well. The loop
may prevent proper expression of both σ 54 and σ 70 promoters.
253
CHAPTER 5
GENETIC ANALYSIS OF THE HRPX PAS-KINASE AND PARTIAL
CHARACTERIZATION OF RECOMBINANT HRPX PROTEINS AND
PROTEIN FRAGMENTS
5.1 INTRODUCTION
In bacteria, most adaptive responses to changes in the environment are, directly or indirectly, mediated by two-component signal transduction systems (Stock et al., 2000). These regulatory systems are composed of a sensor kinase and a response regulator, which is often a transcription factor. Most sensor kinases are transmembrane proteins, presumably perceiving extracellular signals via the periplasmic portions of the input domain. Some, however, are soluble proteins, in particular when the signal perceived is intracellular. Examples of cytoplasmic sensors are E. coli NtrB and Bradyrhizobium japonicum FixL, which sense nitrogen starvation conditions and cellular oxygen levels respectively (McFarland et al., 1981; Taylor and Zhulin, 1999; Gilles-Gonzalez et al., 1994). In other cases, the sensor may be a transmembrane protein with its sensory
254 input domain located in the cytoplasm. Examples of this are the E. coli
ArcB redox sensor and the Sinorhizobium meliloti FixL (Gilles-Gonzalez
et al., 1995; Gong et al., 1998; Gilles-Gonzalez, 2001). Many two- component systems are involved in the regulation of virulence genes
(Hoch and Silhavy, 1995; Finlay and Falkow, 1997). In only a few cases,
however, is the stimulus perceived by the sensor kinase known. For
example, in Bordetella pertussis, expression of hemoagglutinin and toxins
is under the control of BvgS/BvgA, which transduce signals such as
2- temperature, SO4 ions, and nicotinic acid (Arico et al., 1989). In
Shigella flexneri, osmolarity modulates virulence gene expression via the
EnvZ/OmpR system (Bernardini et al., 1990). In Salmonella enterica, the
PhoQ/PhoP, PmrB/PmrA and RcsC/YojN/RcsB pairs form a multicomponent, interconnected cascade responding to extracellular Mg2+,
Fe3+ concentrations, pH and cell envelope stresses (Fig. 1.7) (Soncini and
Groisman, 1996; Kox et al., 2000; Bijlsma and Groisman, 2003;
Chamnongpol et al., 2003; Mouslim and Groisman, 2003). In
Agrobacterium tumefaciens, phenolic compounds and monosaccharides are perceived by the VirA/VirG system that regulates the vir type IV
secretion apparatus leading to the development of crown gall tumors
(Shaw et al., 1988; Lee et al., 1995).
P. stewartii uses the hrpXY two-component system to activate the hrp/hrc type III secretion system. The HrpX sensor PAS-kinase is required for full activation of hrp regulatory and secretion genes in vitro
255 and for consistent infection of corn seedlings (Merighi et al., 2003). In
contrast, the ortholog protein in Erwinia amylovora and Pantoea agglomerans subsp. gypsophiliae is not required for HR elicitation and virulence (Wei et al., 2000b; Mor et al., 2001).
The input region of HrpX contains two tandem PAS sensory domains.
These are cytoplasmic modules of sensor kinases that are often implicated in perceiving a wide range of intracellular stimuli in bacteria, including oxygen levels (via heme prosthetic groups; reviewed in Perutz et al.,
1999), redox conditions (via FAD, FMN, and possibly MK and Qo, binding; Repik et al., 2000; Georgellis et al., 2001; Bock and Gross,
2002), energy levels (via ATP/ADP binding) (Stephenson and Hoch,
2001), and light (Cheng et al., 2002). In sensors with multiple PAS domains (Stephenson and Hoch, 2001; Wang et al., 2001), these motifs are also involved in intermolecular protein-protein interactions and probably the two functions are often overlapping. From previous studies, we know that the expression of genes directly or indirectly controlled by hrpXY respond to environmental variables, such as pH, osmolarity, nitrogen starvation and nicotinic acid (Merighi et al., 2003), but whether
HrpX is actually perceiving any of these stimuli is not known. Similarly, the role of the two PAS domains in the input region of HrpX has not been determined in any of the erwinias.
In this chapter, we describe initial experiments to define a role for the
HrpX PAS domains using genetic and biochemical approaches. Plasmid
256 clones with in-frame deletions of hrpX were constructed for expression in
P. stewartii. The phenotypes conferred by these plasmids in ∆hrpX strains
or the phenotypes of mutants constructed by allelic exchange of these mutations are described. Preliminary results for the purification of the sensor kinase protein and of the tandem PAS domains are presented as
well as the construction of several recombinant plasmids that can be used
to purify HrpX and its domains in future studies. We also used the effect
of HrpY[D57N] overexpression, which overrides the need for
phosphorylation (as described in Chapter 2), to define which classes of
signals may be perceived by HrpX. This strategy was chosen because we
do not have a constitutively active, mutant form of HrpX.
5.2 MATERIALS AND METHODS
5.2.1 Media and Strains and general microbial genetic techniques
E. coli and P. stewartii strains and plasmids used in this study are listed
in Table 5.1. Luria-Bertani (LB) broth and agar (Ausubel et al. 1987) were used for strain maintenance and for growth of P. stewartii under
Hrp-repressing conditions. Fusaric acid-containing tetracycline sensitive selection (TSS) medium was prepared as described by Metcalf et al.
(1996). To induce hrp genes in P. stewartii strains, overnight LB broth
cultures were washed twice in inducing medium (IM5.5; Frederick et al.
2001) prepared without casaminoacids and with only 10 mM sucrose
(IM5.5a), inoculated into 2 to 5 ml of the same medium at A600nm ~ 0.05,
257 grown for 16 h and adjusted to a final A600nm of ca. 0.5. Standard IM5.5,
as described in Chapter 2, was used for GUS assays with the hrpX
mutants. In the experiments where chemicals were tested for their activity
on hrp expression, utmost care was taken to assure that the various media
had comparable osmolarity. Carbon and nitrogen sources were used at the
concentrations reported in the respective figure legends. When practical,
the van ’t Hoff law was used to calculate the approximate osmotic
pressure of the various media. HEPES or Tris-HCl buffers were used in
place of MES for the neutral and basic pH ranges, respectively. Liquid
cultures were grown in flasks or tubes shaken at 200 rpm at 37°C for E.
coli or 29°C for P. stewartii. When appropriate, antibiotics were supplied
at the following concentrations: ampicillin, 200 µg ml-1; kanamycin, 50
µg ml-1; and tetracycline, 20 µg ml-1 (or 15 µg ml-1 for selection of single copy Tetr cointegrates). In IM liquid media the concentration of
antibiotics was reduced by half.
5.2.2 Protein modeling
Modeling of the secondary structure of HrpY was performed with the
PSI-PRED modeling server (McGuffin et al., 2000). Threading of portions of the HrpX amino acid sequence into known crystal structures was performed with mGenTHREADER. Both servers are available at http://bioinf.cs.ucl.ac.uk/. The conserved residues in Bradyrhizobium
japonicum FixL crystal structure (PDB accession 1DRM) were identified 258 with mGenTHREADER and rendered as ball-and-stick atoms in secondary
structure schematics. Molecular models were visualized and rendered with
Acelrys ViewerLight v. 5.0 (Acelrys). Multiple sequence alignments were
performed with Clustal X. Phylogenetic trees were designed with
TreeView and the branches were resampled by 1000 cycles of
bootstrapping.
5.2.3 Construction of strains with in-frame deletions in various hrpX
domains
Unmarked in-frame deletions spanning various portions of the input domain of hrpX were constructed in vitro by PCR as previously described
(Merighi et al., 2003). The PCR fragments and positions of the primers are shown in Fig. 5.1. For the PCR reactions, plasmid pMM58 was used as the template DNA and Pfu as the DNA polymerase (Clontech). The
deletion spanning amino acids 15 to 198 (eliminating both PAS1 and only
part of the PAS2 core) was constructed by amplifying flanking DNA
sequences with primers XdX4AB+BdX5B (fragment C) and primers
BdX1+EdX1 (fragment B) (Table 5.2 and Fig. 5.1). A second ∆hrpX
allele had a deletion between residues 15 and 270, spanning PAS1, PAS2
and part of the linker and this was constructed by amplifying flanking
DNA (fragments D and B) with primers XdX4+BdX5A and primers
BdX1+EdX1, respectively. PCR fragments were digested with the
259 restriction enzymes shown in Fig. 5.1 and ligated into pBluescriptKS
(Stratagene) as contiguous EcoRI-BamHI and XbaI-BamHI inserts to produce plasmids pMM243 (∆hrpX15-198) and pMM242 (∆hrpX15-270), respectively. Deletions of the individual PAS domains were constructed by SOE-PCR. To construct the ∆hrpX15-130 allele, primers
dXPASR2+dXPAS1F2 and primers dXPAS1R1+dXPASF1 were used to
amplify fragments E and F (Fig. 5.1). The two PCR products were gel
purified, mixed in a 1:1 molar ratio, melted, annealed and then used as
templates for a second round of PCR with primers dXPASR2+dXPASF1.
The recombinant PCR fragment “E+F” was gel purified and cloned into
pBluescriptSK as BamHI+EcoRI insert to produce pMM283. A similar
strategy was used to construct the ∆hrpX15-130 allele, with the difference that primers dPAS2F2 and dPAS2R1 were used to delimit the deletion
(Fig. 5.1). The recombinant fragment G+H was cloned in pBluescriptSK
(Stratagene) to produce pMM287. All pBluescript recombinant plasmids were verified by sequencing both strands before any further manipulation.
To generate suicide plasmids for allelic exchange, the inserts in pMM242, pMM243, pMM283 and pMM287 were subcloned as XhoI+SacI fragments and ligated into pLD55, to create plasmids pMM249, pMM247, pMM324 and pMM327, respectively. The suicide plasmids were used to generate unmarked mutations in the P. stewartii DC283 chromosome following the
“Buchner selection” protocol described in Chapter 2. The mutant strains
260 were named MM284 (DC283 ∆hrpX15-270), MM266 (DC283 ∆hrpX15-198),
MM360 (DC283 ∆hrpX15-130), and MM348 (DC283 ∆hrpX143-259) (Fig.
5.2). All mutants were verified by PCR and Southern blot analysis.
5.2.4 Construction of plasmids for genetic complementation tests
Plasmids containing hrpX alleles expressed from their own promoters
were constructed by cloning SOE-PCR fragments amplified from plasmids
pMM283 and pMM287 or by subcloning restriction fragments from
pMM58. To construct a plasmid expressing hrpX from its own promoter,
pMM58, which contains the entire hrp regulatory region, was digested with PshAI and FspI and a resulting 2.1 kb kb fragment was ligated into pBKS at the EcoRV site to produce plasmid pDM2890. This insert in this plasmid was then excised by digestion with EcoRI and HindIIII and recloned into pRK415 such that hrpX was downstream of the Plac promoter in the vector; the resulting plasmid was named pMM389 (or pDM2893). To construct plasmids expressing hrpX alleles missing just one PAS domain (i.e. ∆hrpX15-130 and ∆hrpX143-259), inserts from plasmids pMM283 and pMM287 were excised with EcoRI and BamHI and then ligated into pRK415 to produce pMM378 and pMM380, respectively.
261 5.2.5. Construction of plasmids for expression and purification of
HrpX
The entire hrpX ORF and various portions of it were cloned into several protein expression vectors to produce recombinant proteins with tags to facilitate their purification. Vectors pET28b was used to create C-terminal poly-histidine fusions, while pET15b and pQE30 were used for N-terminal poly-histidine fusions. N-terminal fusions to MBP were created with plasmid pMALc2X , while fusions to GST were constructed in plasmids pGEX-2T and pET41b. C-terminal Chitin Binding-Intein domain fusions were made with vector pTYB1. All constructs were produced by 15-20 cycles of PCR using Pfu DNA polymerase and pMM58 as DNA template.
The cloned inserts were verified by sequencing both strands. The full length ORF was amplified by PCR with primers ETXF1+ETXR (for vector pET15b), primers XBAMF(+1)+ETXR (for vectors pQE30 and pGEX-2T) or primers XpMALF+XpMALR (for vector pMALc2X) (Table 5.2).
Products were gel purified, digested with BamHI and ligated in to plasmids pET15b, pQE30, pMALc2X and pGEX-2T to produce plasmids pMM228, pMM377, pMM315 and pMM371, respectively (Fig. 5.3).
Primers XcHISF2+XcHISR2 (Table 5.2) were used to amplify the hrpX
ORF for cloning into pET28b; the products were gel purified and cloned
into the vector as XhoI/NcoI restriction fragments to generate plasmid pMM367 (Fig. 5.3). Finally, to construct a plasmid expressing a HrpX-
Sce-CB fusion, the hrpX ORF was amplified with primers
262 XNHEF+XNHER (Table 5.2) and ligated as a XhoI/NdeI restriction
fragment into vector pTYB1 to produce plasmid pMM362 (Fig. 5.3). The
insert from pMM228 was also cloned into pEXT20 as an XbaI-HindIII
fragment to produce plasmid pMM319, which expresses hrpX from a Ptac promoter.
The transmitter, input and individual PAS domains were cloned in pET15b as BamHI PCR fragments for expression as poly-His tagged proteins (Fig. 5.3). Primers ETXF1 and ETPAS2R (Table 5.2) were used to amplify the input domain-encoding DNA from amino acids 1 to 485 and produce plasmid pMM257, primers ETXF1 and ETPAS1R amplified the
PAS1 domain (amino acids 1 to 142) to produce pMM251, primers
ETPAS2F and ETPAS2R were employed to produce a fragment expressing the PAS2 domain (amino acids 136 to 276) in plasmid pMM280, and finally primers ETXR1 and ETXF2 amplified the transmitter domain from amino acids 277 to 485 in plasmid pMM233. All inserts were sequenced from both directions to check for polymerase errors. The inserts from pMM257 and pMM251 were also subcloned into the BamHI site of pET41b in order to express N-terminal GST-His6 -S-tag protein fusions
from plasmids pMM270 and pMM265, respectively. The input domain was
also cloned into pTYB1, as an NheI/XhoI PCR fragment amplified with
primers XNHEF and PAS2-Xho-R, to produce plasmid pMM363, and into
pGEX-2T to produce a simple GST-PAS1-2 fusion expressed from plasmid
263 pMM374. For the latter, primers XBAMF(1+) and ET-PAS2R were used to
amplify a PCR product, which was ligated into the vector as a BamHI
restriction fragment.
2+ 5.2.6 Overexpression and purification of His6 -HrpX protein by Ni -
NTA affinity chromatography
Plasmid pMM228 was transformed by electroporation into E. coli
BL21(DE3) carrying pLysS. Several independent Apr Cmr isolates were
stored at –20 and -80°C in 40% glycerol, 10 mM MgSO4 , 50 mM Tris-HCl pH 7. For protein expression, bacteria were grown overnight in 5 ml LB broth with antibiotic selection. Two 2-l flasks containing 700 ml LB broth were inoculated with 1.5 ml of overnight culture. The flask was shaken (200 rpm) at 37°C until the culture reached A600=0.57 (measured
in a B&L Spectronic 20 colorimeter with 13 mm x 100 mm tubes). IPTG
was added to a final concentration of 1 mM and the culture was grown at
15°C for 15.5 h to a final A600=1.7. Cells were harvested by
centrifugation in a Sorval GSA rotor at 7000 rpm for 20 min at 4°C.
About 12 g (wet weight) of cells were resuspended in 46 ml ice-cold lysis
buffer (10 mM imidazole, 500 mM NaCl, 5 mM MgCl2 , 10 mM β-
mercaptoethanol, 0.5 mM PMSF, 1 µg ml-1 leupeptin 0.1% NP-40, 5%
glycerol, 20 mM Tris-HCl pH 7.9 at 4°C). The cell suspension was
incubated in 0.5 mg ml-1 lysozyme for 60 min on ice and then passed three
times through a 40-K French pressure cell (Thermo IEC) at 10,000 psi 264 adding 1 mM PMSF after the first passage. The lysate was centrifuged
twice at 17,500 x g for 45 min at 4°C. The supernatant was incubated with
4 µg ml-1 pancreatic DNase I (Sigma) for 15 min at room temperature and
then it was centrifuged at 17,000 x g for 10 min. IMAC column
chromatography of the supernatant was performed at 4°C using a 1.5 ml
glass column (I.D. = 1 cm) packed with 3 ml of Ni2+-NTA agarose slurry
(Novagen) and equilibrated with binding buffer (10 mM imidazole, 500
mM NaCl, 5 mM MgCl2 , 5 mM β-mercaptoethanol, 20 mM Tris-HCl, pH
8.0 at 4°C). The average flow rate was 0.3 ml min-1. The column was
washed with eight column-volumes of binding buffer, ten column-volumes
of washing buffer (40 mM imidazole, 500 mM NaCl, 5 mM MgCl2 , 20 mM
Tris-HCl pH 8.0 at 4°C) and five volumes of elution buffer A (80 mM imidazole, 500 mM NaCl, 5 mM MgCl2 , 20 mM Tris-HCl pH 8.0 at 4°C).
The target His6 -HrpX proteins were eluted with 5 ml 500 mM imidazole
buffer (500 mM imidazole, 500 mM NaCl, 5 mM MgCl2 , 20 mM Tris-HCl
pH 8.0 at 4°C) collecting 1-ml fractions. The target protein migrated in
SDS-PAGE as a 63 kDa band instead of 58 kDa as expected. Fractions
containing higher concentrations of the 63Kda protein were pooled and
either dialyzed overnight against 2 l of storage buffer (100 mM KCl, 1
mM MgCl2, 1 mM DTT, 0.1 mM EDTA, 10% glycerol, 20 mM Tris-HCl
pH7.9) or buffer-exchanged by repeated ultrafiltration in a MicroCon-10
spin filter (Amicon). The semipurified protein preparation was stored at –
265 80°C. The degree of purification was estimated by Coomassie Brilliant
Blue staining (Laemmli, 1970) and the fractions were analyzed by
Western blot using anti-His6 monoclonal antibodies (Novagen) following
the manufacturer’s instructions.
5.2.7 Overexpression and purification of MBP-HrpX protein by
amylose affinity chromatography
Plasmid pMM315 was transformed by electroporation into E. coli strains
DH10B, TB1 and ER2507. For protein expression, bacteria were grown
overnight in 10 ml LB broth with antibiotic selection and 0.2% glucose.
One 2.8-l flask containing 1000 ml LB broth, 200 µg ml-1 ampicillin,
0.2% glucose was inoculated with 1.5 ml of overnight culture. The flask
was shaken (200 rpm) at 37°C until the culture reached A600=0.65. IPTG
was added to a final concentration of 0.3 mM and the culture was grown
at 15°C with shaking for 16 hr. Cells were harvested by centrifugation in
a Sorval GSA rotor at 8000 rpm for 20 min at 4°C. About 12 g (wet
weight) cells were then resuspended in 25 ml of ice-cold column buffer
(CB; 2 mM NaCl, 10 mM β-mercaptoethanol, 1 mM PMSF, 1 µg ml-1 pepstatin, 20 mM Tris-HCl, pH 7.4 at 4°C). The cell suspension was incubated in 0.5 mg ml-1 lysozyme for 20 min on ice and then passed two
times through a 40-K French pressure cell (Thermo IEC) at 10,000 psi
adding 0.5 mM PMSF after the first passage.The lysate was centrifuged
266 twice at 20,000 x g for 45 min at 4°C. The supernatant was sonicated to reduce the viscosity and diluted by adding 100 ml of CB. Column chromatography was performed at 4°C. The supernatant was then applied to a glass column (I.D.= 1 cm) packed using 6 ml of an 80% amylose slurry (Novagen) and pre-equilibrated with 40 ml of column buffer. The average flow rate was 0.3 ml min-1. The column was washed with 12 column-volumes of CB buffer. The MBP-HrpX protein was eluted with four column volumes of elution buffer (10 mM maltose in column buffer) collecting 1-ml fractions. Fractions containing the expected 100-kDa band were pooled and concentrated by ultrafiltration in a Vivaspin 15R device.
The protein preparation was proteolyzed with 0.25% factor Xa for 24 h at
4°C followed by 24 h incubation at room temperature. The protein mixture was dialyzed in buffer A (20 mM Tris-HCl pH8, 25 mM NaCl, 10 mM β- mercaptoethanol) for 24 h and fractionated by ion-exchange chromatography. A linear gradient starting with 100% buffer A and 0% buffer B (20 mM Tris-HCl pH8, 750 mM NaCl, 10 mM β- mercaptoethanol) and ending with 100% buffer B was used. Fractions containing the 56-kDa target protein were pooled and concentrated by ultrafiltration. The degree of purification was estimated by Coomassie
Brilliant Blue staining (Laemmli, 1970).
267 5.2.8 Overexpression and purification of GST-PAS1-2HrpX protein by
glutathione affinity chromatography
For expression of the GST-PAS1-2HrpX protein, E. coli DH10B
(pMM374) was grown overnight in 10 ml LB broth with antibiotic
selection. One 1-l flask containing 100 ml LB broth, 200 µg ml–1 ampicillin, 0.2% glucose was inoculated with 1 ml of spun and washed overnight culture. The flask was shaken (200 rpm) at 37°C until the culture reached A600=0.6. IPTG was added to a final concentration of 1
mM and the culture was grown with shaking at 37°C for 4 hr. Cells were
harvested by centrifugation in a Sorval GSA rotor at 8000 rpm for 20 min
at 4°C (wet weight 12 g) and then resuspended in 5 ml of ice-cold PBS
buffer (1X phosphate buffer saline pH 7.4). The cell suspension was
incubated in 0.5 mg ml-1 lysozyme for 20 min on ice and sonicated by 10
bursts of 10 s each at 20% duty, adding 1 mM PMSF after the first burst.
The lysate was centrifuged twice at 20,000 x g for 45 min at 4°C. Column
chromatography was performed at 4°C. The supernatant was applied to a glass column (I.D. = 1 cm) packed with 3 ml of a 50% GST-BindTM resin slurry (Novagen) and pre-equilibrated with 15 ml of PBS. The column was washed with 12 column-volumes of PBS. The target GST-HrpX protein was eluted with three column-volumes of elution buffer (50 mM Tris-HCl pH 8.0, 10 mM reduced glutathione) collecting 0.5-ml fractions. Fractions containing the expected ca. 80-kDa protein were pooled and concentrated by ultrafiltration in a Vivaspin 15R device. 268 5.2.9 Mass spectrometry protein fingerprinting
Protein samples were analyzed by nano-liquid chromatography tandem
mass spectrometry at the OSU CCIC Proteomics and Mass Spectrometry
Facility. Proteins were separated by SDS-PAGE (10% T; mono to bis-
acrylamide ratio = 30:0.8 ), fixed in 5% acetic acid in ethanol followed by
standard Coomassie staining and destaining. Protein bands were excised
and stored in 5% acetic acid at 4°C. An excised gel fragment without
visible proteins was used for the blank control run. The samples were
cleaved by in-gel trypsin digestion. Tryptic fragments were characterized
by MS/MS fragmentation. Data were analyzed using the Mascot server by
comparison to the Genbank protein database. The sequence of 46% of
HrpX was determined followed this procedure.
5.2.10 In vitro autophosphorylation of HrpX and phosphotransfer
reaction
For autophosphorylation of HrpX, 15 µl of a mixture containing 30 µM
HrpX, 0.22 µM [γ32P]ATP (3000 Ci/mmol; NEN) and 0.1 mM cold ATP in
phosphorylation buffer (125 mM potassium glutamate, 125 mM HEPES pH
8.0, 5 mM MgCl2 , 0.1 mM DTT, 5% glycerol) was incubated at 25°C for up to 60 min. Reactions were terminated with 5 µl of 4X stop/sample buffer (125 mM Tris-HCl pH 6.8, 8 mM EDTA, 4% SDS, 8% β-mercapto- ethanol, 20% glycerol, 0.02% bromophenol blue). For transphosphorylation reactions, HrpX was allowed to autophosphorylate 269 for 20 min as described above and then 40 µM HrpY was added. Reactions
were terminated at intervals using 4X stop buffer. Proteins were separated
by SDS-PAGE (12% monomer gel), and visualized by Coomassie Brilliant
Blue staining (Laemmli, 1970). Gels were dried under vacuum at 60°C for
30 min. The radiolabeled proteins were visualized by autoradiography at –
80°C using Kodak MS film.
5.2.11 UV-crosslinking of the HrpX input domain with [γ 32P]ATP
Direct photo-affinity labeling of the PAS domains with [γ32P]ATP was performed following the protocol of Stephenson et al. (2001). In brief, 50 ng of recombinant protein was incubated in 50 µl reaction mixtures containing 50 mM HEPES pH 7.5, 5 mM MgCl2 , 0.5% acetone and 2 µM
[γ32P]ATP (80 µCi) for 30 min on ice in the dark. A 40 µl aliquot was
placed in an open Eppendorf tube incubated on ice and exposed to 254 nm
light from a UV cross-linker device (Stratagene) for 15, 30, 45 and 60 min
at 125 J s-1 at a distance of 5 cm from the light. The reactions were
stopped by boiling 10 µl of each mixture for 3 min in sample buffer and
then separating the proteins in Laemmli gels followed by autoradiography.
5.2.12 Pathogenicity tests
Pathogenicity tests were performed by inoculating the whorls of 5 to 6-
day-old sweet corn seedlings (Zea mays var. saccharata cv. “Seneca
Horizon”) as previously described (Coplin et al. 1986). The plants were 270 held in growth chambers at 29°C (photoperiod 16 hr, 15000 lux, relative humidity 99%). Each corn whorl was inoculated with 50 µl of ca. A600nm=
0.1 suspensions of P. stewartii cells in 10 mM potassium phosphate buffer
pH6.8, 0.2% Tween 40. After 3-4 days, disease severity was rated using a
0 to 3 scale (0= no symptoms, 1=scattered small lesions, 2 = numerous lesions and 3 = extensive lesions that remained water-soaked with ooze forming on leaf surfaces).
5.3 RESULTS
5.3.1 The input region of P. stewartii HrpX is formed by two PAS
domains with low similarity between them
In order to rationally create a subdomain deletion in hrpX, we decided
to study in more detail the primary and secondary structure of HrpX. The
amino acid sequence of HrpX was initially analyzed by PSI-BLAST and
CDD searches at the NCBI web site to define potential orthologs and
identify functional domains. The closest orthologs in the erwinias and two
paralogs (hypothesized to be orthologs) in the genus Pseudomonas are
described in Chapter 2. Here their sequences are compared by ClustalX.
Branches of the derived unrooted trees were bootstrapped using the
neighbor-joining method (1000 resamplings). The resulting tree (Fig. 5.5)
shows phylogenetic relationships consistent with the taxonomic position
of the species.
271 Two PAS domains comprised 90% of the input region of the sensor kinase. The first PAS domain (PAS1) spanned residues 16 to 130 and the second domain (PAS2) was located between residues 142 to 259. The C- terminal region of PAS domains, which contains the β-scaffold, is usually poorly conserved, so these coordinates have a higher margin of error in that position. The two domains were 21.2% identical to each other (28% similarity). Based on the CDD search results, the PAS1 core was 100% identical to the consensus motif Pfam00989 (E=1x10-5), whereas PAS2
core was only 89.2% identical to it (E=0.002). Pfam00989 contains only
the highly conserved PAS core, not the helical connector and the β-
scaffold, but similar results were obtained from comparisons with
consensus motif Cd00130.1, which also contains the C-terminal portion of
the PAS domain. The secondary structure of HrpX was predicted using
PSIPRED. The PAS1 domain has a typical α/β fold (topology:
AβBβCαDαEαFαGβHβIβ). Two independent PSI-PRED analyses
predicted that the short linker between PAS1 and PAS2 may fold either as
a β-strand/loop (Fig. 5.6) or as a helix (not shown), but the statistical
confidence was only moderate in both analyses. PAS2 appears to be
missing the Eα helix and contains a slightly longer Dα helix. The pocket
between the Eα and Fα helices and the Gβ, Hβ and Iβ strands in FixL and
Phy3 PAS domains were shown by NMR and X-ray crystallography to
form the ligand binding site (Amezcua et al., 2002). The differences at
this site between FixL and HrpX could reflect the fact that different 272 ligands are bound by the kinase. The HrpX PAS core domains were compared to other known PAS domains in an attempt to predict what ligand(s) might interact with HrpX. A CDD search produced a list of 50 domains most similar to PAS1 out of the many thousands found in
GenBank. It included, in order of decreasing similarity, Rhizobium and
Bradyrhizobium FixL (1EW0 and 1DRM), Rhizobium NodV (gi 128494),
E. coli NtrB (gi 128594), Bacillus subtilis KinC and KinA (gi 729901, gi
3041695), the Neurospora crassa protein WC1 (gi 2494692), and E. coli
NtrY (gi 585587) and ArcB (gi 1168485). The corresponding list using
PAS2 as a query included TraJ (gi 136225), PhoR (gi 1172485), NtrB (gi
128597) and ArcB (gi 1168485). The associated ligands are heme and oxygen for FixL and ATP for KinA; ArcB is believed to bind the respiratory chain cofactors MK3 and Q0 . The phylogenetic trees in Fig.
5.7 show the relative genetic distances between these PAS domains.
Unfortunately, this tree again demonstrates that, based on similarity alone, it is impossible to predict what ligand is bound by any given PAS domain, since WCI, FixL and KinA PAS modules cluster together but bind three very different molecules (FAD, heme, and ATP, respectively).
The threading of the HrpX PAS1 into the FixL PAS backbone crystal structure (1DRM) was performed with mGenTHREADER (Fig. 5.8) and the corresponding three-dimensional model is shown in Fig. 5.9.
Conserved residues are shown as ball-and-sticks. Automated homology modeling with the ProMod algorithm at the SwissModel server failed due
273 to a low level of identity (<25%) to the reference structure, and structural alignments with high RSM (~19Å) were obtained (not shown). This information was helpful in designing in frame deletions of HrpX that did not alter any secondary structure elements that might cause misfolding the mutant protein. These secondary and tertiary structural models may be a useful reference for future studies aimed at engineering mutations in the active site of the PAS structure that will help to identify the ligands.
5.3.2 The two PAS domains of HrpX play different roles in modulating its activity
In frame deletions in the hrpX gene were engineered in vitro to produce plasmids for complementation experiments and constructing chromosomal mutations in P. stewartii. The deletions were designed to preserve the secondary structural elements identified in Fig. 5.6 and minimize the risk of producing locally misfolded proteins. Deletion of the entire hrpX gene in strain DM2790 ∆hrpX17-414 is described in Chapter 2. When inoculated into corn whorls, the virulence of this mutant was only slightly reduced and its watersoaking severity rating was 2.0-2.5 as compared to 3.0 for the wild-type strain (Fig. 5.10). Deletion of both PAS domains in strains
MM266 ∆hrpX15-198 and MM284 ∆hrpX15-270 or just PAS2 in strain
MM360 ∆hrpX143-259 reduced the severity rating to about 2.0, which corresponds to numerous lesions but not collapse of entire leaves. In contrast, deletion of just the PAS1 domain in strain MM348 ∆hrpX15-130 274 strongly reduced virulence to a level below 1.0 (i.e. only a few small,
scattered lesions) (Fig. 5.10). All of the mutants were complemented to
full virulence by pMM389 hrpX+. Interestingly, strain MM360 (∆PAS2)
was fully complemented by pMM380 (∆PAS1), and, vice versa, strain
MM348 (∆PAS1) was complemented by pMM378 (∆PAS2). Intracistronic
complementation suggests that the HrpX protein may dimerize as shown
for other prokaryotic sensor kinases. These data indicate the critical
importance of the PAS1 domain for proper signal transduction in vivo. On
the other hand, PAS2, although required for full virulence, appears only
to cooperate with PAS1. In other words, PAS1 is necessary but not
sufficient for full HrpX activity. Expression of pMM380 in the wild-type
background did not affect virulence (not shown), ruling out negative
trans-dominance of the ∆PAS1 allele. This is somewhat unexpected, because the same allele appears to reduce the virulence of the strain
DM2790 ∆hrpX17-414. Such results imply an active suppression by the
HrpX∆PAS1 protein of the cellular cross-talk to HrpY occurring in this
genetic background (Merighi et al., 2003), while the same mutant protein
appears unable to compete with wild type HrpX.
The expression of a representative hrp secretion gene (hrpJ-uidA in
pDM2531) in the above hrpX mutants was not always correlated with their
virulence (Fig. 5.11). We observed that while the deletion of the entire
hrpX gene in DM2790 and the ∆PAS1-PAS2 deletion in mutant MM284
reduced hrp expression by 90% from the wild-type levels, these mutants
275 still produced 70% as much watersoaking as the wild-type strain. These relative levels of expression were comparable to those observed in planta by epifluorescence microscopy of the same strains carrying hrpJ-gfp fusions (not shown). This implies that the transmitter domain in the
∆(PAS1-PAS2) mutants is not functioning as a kinase, and that, both in planta and in vitro, cross-talk between HrpY and other kinases or small phosphodonors is controlling hrp gene expression in these genetic backgrounds (Chapter 2 and Fig. 5.11). On the other hand, both hrpJ
expression and virulence was drastically reduced in the ∆PAS1 mutant
MM348 (1.8% of wild-type). (Fig. 5.11). In contrast, the ∆PAS2 strain
(MM360) expressed hrp genes in IM5.5 at about 20% of the wild-type level but was only slightly affected in virulence (watersoking score ~2).
In summary, PAS1 seems to be absolutely required for both stimuli perception in vitro and virulence in planta. Therefore, it is possible that this truncated protein is locked in a conformation that suppresses cross- talk (as shown in strain DM2790/pMM380; Fig. 5.10) or/and the functioning of the PAS2 domain. For example, this could happen if it is locked in a phosphatase mode. On the other hand, PAS2 is still required for full activation of the sensor protein. One possibility is that PAS1 and
PAS2 sense separate signals and then act cooperatively to control the kinase domain. Another possibility is that PAS2 might function as an helper of PAS1, by being involved in dimerization of the sensor molecules, as shown for Bacillus KinA.
276 Mutant MM266 does not contain a full deletion of the PAS2 domain and
a small portion of PAC2 motif carrying the regulatory Fα helix-F-G loop is preserved. hrp expression in MM266 is only 1.2% of the wild-type, similarly to the ∆PAS1 strain MM348. However, differently from MM348, virulence was not affected as severely. The reason for these differences is unclear.
5.3.3 HrpX does not transduce pH, osmolarity or many catabolic metabolite-associated signals but it appears to sense stimuli related to
nitrogen metabolism and the Krebs cycle
As shown in Chapter 2, many different environmental signals and/or
growth conditions affect the expression of hrp genes. To start dissecting
which of these stimuli are perceived by the HrpX/HrpY signal
transduction system, we determined which repressing conditions could be
overridden by overexpression of HrpY and ergo might be affecting HrpY
activation. This strategy assumes that other conditions operating directly
on hrpS would not affect HrpY overexpression. To do this we exploited
the observation that ectopic expression of the hrpY[D57N] allele in
pMM118 activates hrpS expression independently of any phosphorylation
event catalyzed by HrpX. The use of the mutant hrpY[D57N] gene instead
of wild-type hrpY eliminates the risk that phosphorylation or
dephosphorylation of HrpY might confuse (dampen or stimulate) its
effects on hrpS. P. stewartii strain DM733 hrpY::Tn5 hrpS-lacZ was used 277 as a recipient for pMM118 (Fig. 5.12). DM733 has very low basal β- galactosidase activity and was used as negative control in all experiments
(data not shown). Strain MEX1 hrpS-lacZ was used a positive control and enzyme activity measurements were corrected for the basal β- galactosidase activity in DC283. Bacteria were grown in IM5.5A with one or two chemical variables changed at a time, β-galactosidase activity was measured and data were compared to the expression of each strain in
IM5.5A (data in Fig. 5.12 expressed as a percentage of activity in
IM5.5A).
Elevated osmolarity due to potassium phosphate or sodium chloride caused repression of hrpS in the MEX1 hrpS-lacZ control strain and overexpression of hrpY[D57N] did not override this effect (Fig. 5.12).
Likewise, the reduced expression of hrpS in media with high pH,
asparagine as a sole source of C and N, nicotinic acid at 1 mM or peptone
was not increased by hrpY overexpression. Curiously, at higher population
density, repression of hrpS in cells grown in IM5.5 with asparagine as the
sole C/N source was overcome by pMM118 in one experiment. The
expression hrpS-lacZ fusion was strongly reduced by growth in IM5.5
with tryptone, alone and in combination with sucrose (4.5-fold repression)
and overexpression of hrpY[D57N] restored hrpS transcription. In
contrast, the low hrpS-lacZ expression observed in LB broth, which also
contains tryptone, was not increased by pMM118. Presumably, this is
because LB broth also contains high salt concentrations (0.5%). The 278 downregulation of hrpS by growth in IM with citric acid or succinate as sole C source was variable and ranged from 40-60%, compared to the
IM5.5 control, but hrpS transcription was always fully restored by
pMM118. Other respiratory catabolites, such as pyruvate, aspartate and
glutamate, did not have any significant effect on hrpS expression (not
shown).
5.3.4 Expression and Purification of recombinant HrpX proteins
Several types of vectors and polypeptide fusions were used to engineer
versions of HrpX that could be purified by affinity chromatography. This
was necessary because initial experiments with a His6 -HrpX fusion
indicated that we might encounter solubility problems. Consequently,
various combinations of tags, promoters of different strength and fusions
to various locations in HrpX were constructed so that we could be ready
to switch expression construct/purification chemistry if required.
Initially, His6 -HrpX was overexpressed from pMM228, which uses the
T7lac promoter in a pET-15b vector. A PCR insert was engineered to
maintain the correct translation frame with a vector-encoded N-terminal
polyhistidine tag (ca. 2 kDa in size). The his6 -hrpX gene was also
subcloned from pMM228 into pEXT20 as a HindIII+XbaI fragment for
expression from Ptac (plasmid pMM319). Two additional His6 -tagged versions of this gene were constructed in pET28b and pQE30 (Fig. 5.3) to express proteins with minimal size tags in C- and N-terminal positions
279 respectively. Other tags were also engineered in order to increase the recombinant protein solubility and yield. In particular, N-terminal genes expressing the maltose binding protein, encoded by malE (MBP), and the
glutathione S-transferase polypeptide (GST) were translationally fused to
hrpX and expressed from pMM315 and pMM371, respectively. Finally, a
C-terminal fusion of hrpX to the Intein-Chitin binding domain encoded in
plasmid pTYB1 was engineered in plasmid pMM362. This last construct is
expected to have increased solubility, comparable to the pMM315 MBP-
HrpX fusions, but the removal of the tag is performed in vitro using an
autocatalytic mechanism triggered by the intein domain in presence of
high concentrations of reductants, such as DTT. Most of these constructs
were prepared recently as alternatives to pMM228 and pMM315, which
gave mixed results, as shown below.
Induction of the T7lac promoter of plasmid pMM228 in E. coli
BL21(DE3)(pLysS) by IPTG resulted in the production of a ca. 63-kDa
polypeptide visible in total cell protein lysates, but not present in
uninduced cells (Fig. 5.13). Fig. 5.14 shows the purification steps by immobilized metal affinity chromatography (IMAC) of the poly(His) tagged HrpX proteins using step-elution gradients. Highly purified His6 -
HrpX was concentrated with a VivaSpin 15R ultrafiltration device and the
purified fraction dialyzed against storage buffer is shown in Fig. 5.14 -B.
The purity of the samples was assessed by densitometric analysis of
Coomassie-stained SDS-PAGE gels and by nano-LC-MS/MS spectrometry.
280 In many cases, the purity indicated by SDS-PAGE was >80%, but a
second band of about 30 kDa always copurified with the 63-kDa band,
which is close to the expected size of 58 kDa. Small alterations of
electrophoretic mobility in Laemmli gels are observed for proteins with
hydrophobic domains or with a high proportion of positively charged
amino acids. Western blot analysis using anti-His monoclonal antibodies
confirmed both the 63-kDa and the 30-kDa bands were His-tagged
proteins (Fig. 5.14, C). This suggests that the 30 kDa protein may be a
proteolytic fragment from HrpX (probably corresponding to the PAS1-2
input domain; amino acids 1 to 260, ca. 30 kDa) released during the
purification procedure. This may be due to the fact that metalloprotease
inhibitors were not added because they would interfere with the
immobilized metal affinity chromatography. The average yield was of
0.25 mg of purified soluble protein per liter of culture.
To determine the sequence identity of the 63-kDa polypeptide, the band was excised from an SDS-PAGE gel and subjected to tryptic digestion.
The carbamidomethyl-modified peptides were separated by liquid chromatography and individually fragmented in a MS/MS detector (Fig.
5.15). Thirty peptides, corresponding to 46% of the sequence of the protein, were isolated and characterized. Peptide sequences were matched by Mascott searches to the GeBank accession gi 9885631 corresponding to
HrpX. This His6 -HrpX preparation was used for the autophosphorylation and transphosphorylation experiments described below. The major
281 problem with the recombinant HrpX expressed from pMM228 was its low solubility and yield and the difficulty of obtaining reproducible results between purification batches. This limited the number and complexity of experiments we conducted with His6 -HrpX from pMM228 and forced us to use different His6 -tagged constructs. We attempted various strategies,
including the reduction of the expression level by using weaker Ptac
promoters (plasmid pMM319), the repositioning of the polyHis tag in the
C-terminal position (plasmid pMM367), and the engineering of minimal
polyHis tags (plasmid pMM377). We also tried the expression and
purification of HrpX in P. stewartii DM2790(pMM319), hoping that the
presence of the cognate response regulator might stabilize the
overexpressed protein. None of these approaches helped in solving the
solubility problems.
We then switched to other purification chemistries involving the use of
larger, well structured and soluble recombinant tags. The production of
highly soluble MBP-HrpX recombinant proteins was achieved by
expressing pMM315 in E. coli DH10, ER2507 or TB1. The expected 100-
kDa fusion protein was observed in IPTG-induced samples (Fig. 5.16) and
amylose affinity chromatography was used to purify it. A large amount of
recombinant protein was obtained (ca. 10 mg l-1 of culture). Proteolytic
fragments of ca. 70 kDa were present in spite of the use of a wide-
spectrum protease inhibitor cocktail. We did not attempt to use E. coli
backgrounds with mutations in housekeeping protease systems. The
282 protein preparations eluted from the amylose column were hydrolyzed by factor Xa to release the MBP tag, which was subsequently removed by ion-exchange chromatography (Fig. 5.16). The final yield of soluble tag- less HrpX was 5.6 mg l-1. Removal of the tag was necessary because the
MBP-HrpX fusion protein had extremely weak autophosphorylation activity (not shown), probably due to interference by the N-terminal MBP polypeptide. The caveat was that the removal of the tag resulted in totally inactive kinase preparations
Plasmids pMM371 and pMM362 were only verified for the production of the expected recombinant protein and not tested exhaustively for the purification of soluble recombinant proteins. These are available for future experimentation and should facilitate the preparation of larger amounts of soluble, active HrpX.
5.3.5 Autophosphorylation kinetics and phosphotransfer ability of
HrpX
This section describes functional experiments performed with the only active HrpX preparation obtained. The ability of His6 -HrpX to autophosphorylate was assessed in vitro. The purpose of this experiment was to optimize the conditions for maximal phosphorylation of the sensor kinase so that we could use it to phosphorylate HrpY. In addition, this
283 information will be needed for future studies to test if potential, known
ligand molecules can interact with the PAS domains and affect
autophosphorylation rates in end-point experiments.
Clear autophosphorylation was demonstrated with one preparation of
His6 -HrpX (Fig. 5.17). This was maximal after 60 min of incubation with
0.22 µM of [γ32P]ATP. We were able to use this enzyme preparation to perform a preliminary experiment to test the ability of several known PAS ligands to modulate the rate of autophosphorylation of HrpX. These results indicated that ADP inhibits the reaction, while AMP and menaquinone weakly stimulate it (not shown). A kinetic analysis to determine their effect on the reaction rate was not done for lack of enzyme.
Multiple subsequent attempts to re-purify His6 -HrpX for further
experimentation yielded very little protein and the preparations had no
measurable autokinase activity. Furthermore, the larger amounts of
soluble HrpX obtained from MBP-tagged proteins also showed little or no
autophosphorylation activity (Fig. 5.18). When either form of the protein
was used in transphosphorylation reactions with HrpY, we never observed any clear signals corresponding to HrpY~32P, except in one experiment
using the His6 -HrpX preparation where we observed a radioactive signal
corresponding to a 26-kDa band after 5 to 10 min incubation with
suboptimal concentrations of HrpX (Fig. 5.18). Salmonella BarA/HrpY
reaction mixtures were used as positive controls. The difficulties in
284 obtaining larger amounts of soluble His6 -HrpX halted this line of experiments. Our difficulties in repurifying His6 -HrpX were quite
frustrating. Similar problems of irreproducible purification of soluble
proteins with large hydrophobic domains in general, and PAS-kinases in
particular have been described. These problems are especially exacerbated
by the absence of sufficient amounts of the cognate ligand in cells
overexpressing the recombinant protein (Dr. K. Gardner, pers. comm.).
PAS domains in absence of the cognate ligand tend to compact in an
astructured conformation. To this respect, in two instances we attempted
without success to add ATP and FAD to cleared cell lysates hoping to
increase the protein solubility. On the other hand, we suspect that the
absence of activity for the tag-less MBP-HrpX derivative is due to the
digestion of the preparations at room temperature for 24 h to efficiently release the MBP tag. The kinase may have lost activity during this time.
In the future, it might be better to use highly soluble tags that can be
released more quickly at low temperature (e.g. pMM362).
5.3.6 GST-PAS1-2 does not bind ATP
Some PAS kinases, such as Bacillus KinA (Stephenson et al., 2000), use
ATP to sense internal energy levels in the cell. To test if HrpX may be
sensing ATP levels, we tried to purify subdomains corresponding to the
input region of HrpX. His-tagged recombinant proteins (Fig. 5.13) were
insoluble, and instead of trying to repurify them from the inclusion
285 bodies, we decided to engineer more soluble tags in the C-terminus.
Several other expression plasmids were constructed (Fig. 5.4) and small- scale purifications were attempted. Plasmids pMM270 and pMM265 did not show any upregulated polypeptide upon IPTG induction, possibly because of the instability of the recombinant protein. A soluble recombinant GST-PAS1-2 protein was successfully purified from a small- scale batch of E. coli TB1 (pMM374) (Fig. 5.19A). The recombinant protein spans the whole input domain of HrpX. The protein was soluble, with only a little localizing in inclusion bodies. For the purification, we used a simple affinity column without downstream purification and the protein appear >90% pure in a Coomassie-stained gel. The protein was incubated with a large molar excess of [γ32P]ATP. A clear signal corresponding to the size of the GST-PAS1-2 fragment was not observed
(Fig. 5.19A), so it is unlikely that HrpX binds ATP. As a positive control, we showed that exposure of the reaction mixture to UV light resulted in cross-linking of radiolabeled ATP to background proteins that were not visible in Coomassie-stained gels of the GST-PAS1-2 preparation (Fig.
5.19A). These experiments should be repeated with higher concentrations of more purified protein and with batches of GST-PAS1-2 with the GST tag removed. It is possible that the tag could be interfering with the binding, although this is unlikely due to the small size of the ligand involved.
286
5.4 DISCUSSION
P. stewartii HrpX is a sensor kinase that is important for the
transduction of signals generated by growth in IM5.5, an apoplast
mimicking medium, and for full and consistent virulence on corn
seedlings (Merighi et al., 2003). The input domain of HrpX is composed
of two tandem PAS domains, designated PAS1 and PAS2. In this study,
we found that HrpX PAS1 and PAS2 are not redundant. Their sequence
similarity is low (28% similarity; 21.2% identity) and the effects of
deleting PAS1 or PAS2 are not identical. PAS1 appears to be essential for
signal sensing in IM5.5 (Fig. 5.10) and it is very important for virulence
on corn. In contrast, PAS2 appears to play a secondary role in virulence.
Specifically, the virulence of the ∆PAS1 (∆hrpX15-130) strain was about 1
(“small scattered lesions”), while that of the ∆PAS2 mutant (∆hrpX143-259)
was about 2 (“many lesions”). In this regard, the ∆PAS2 mutant is roughly
comparable to the ∆hrpX17-414 strain (DM2790) (Merighi et al., 2003)
(Fig. 5.10). Thus, PAS2 does appear to be involved in signal sensing in
IM5.5 (Fig. 5.10) but to a much lesser extent than PAS1. It is important to
note that the ∆hrpX15-130 allele does not seem to be negatively dominant,
because it did not reduce virulence in the wild-type background. On the
other hand, its expression in the ∆hrpX17-414 strain seemed to eliminate cross-talk with HrpY, as inferred by a reduction in the virulence of this strain (Fig. 5.10). If the expression of hrp genes in a wild-type strain is 287 affected by the trans-expression of the ∆hrpX15-130 allele is not known.
Finally, intragenic complementation suggests that HrpX molecules can oligomerize.
In other sensor proteins, PAS domains have been shown to control the autophosphorylation activity of the kinase core. Ligand binding to the
PAS pocket leads to structural rearrangements of the Fα-FG-Hβ interaction surface in the PAS fold and modulation of the kinase activity of the catalytic domain (Amezcua et al., 2002). This happens even when purified PAS domain fragments are added to in vitro reactions containing the kinase domain (Rutter et al., 2001). Our genetic data seem to suggest that the two PAS domains in HrpX cooperatively interact to control HrpX autophosphorylation, but that ligand interactions at PAS1 may be dominant over events at PAS2. Consistent with this model is the possibility that PAS2 may work also as a secondary dimerization domain, similar to what has been described for KinA (Wang et al., 2001). We have started to analyze the protein-protein interactions of the individual PAS domains with each other and with the kinase core using a yeast two-hybrid system (Merighi, Ham and Coplin, unpublished), but so far we have not been able to show interactions between the input domain and full-length
HrpX protein fused to CytoTrap tags. The mapping of these interactions may provide insights into the mechanism of regulation of the kinase core by the PAS modules.
288 The nature of the ligand/signal sensed by the PAS domains is of
particular theoretical and practical interest. Very few PAS domain ligands
have been identified under natural conditions. Among those known are 4-
hydroxy-cinnamic acid, FAD, heme, ATP and FMN. A recent NMR screen of 750 organic compounds showed that the hydrophobic core of the hPASK PAS-A domain can specifically bind small molecules (Amezcua
et al., 2002). In this system and several others, ligand binding is
important for the transition between the unstructured state of the
apoprotein and the folded state of the conjugated protein. The tendency
apo-PAS domains to precipitate in the absence of ligand may explain why
we could not obtain soluble His6 -tagged HrpX and PAS proteins, while
addition of highly structured tags, such as MBP or GST, allowed the
purification of reasonable amounts of HrpX or PAS1-2 protein. The low
activity of the purified proteins (especially the MBP-HrpX version), in
addition to thermal inactivation for the reason discussed above, may be
due to loss of the cofactor during purification, especially during dialysis.
Using the only batch of soluble His6 -HrpX we were able to obtain, we
demonstrated rapid autophosphorylation of HrpX, which reached
saturation around 60 minutes from the start of the reaction (Fig. 5.19).
This is comparable to what has been observed for other sensors (Loh et
al., 1997). We do not know if any ligand was copurified with the His6 -
HrpX protein. Absorbance spectral data did not reveal any signals
consistent with the binding of flavin cofactors (not shown). An attempted
289 transphosphorylation experiment with the same protein preparation resulted in a weak signal consistent with the production of HrpY~P, but the negative control was ambiguous due to the presence of background kinases (Fig. 5.18). In future experiments, these problems might be overcome by the use of pMM362 as expression plasmid, which should allow for the expression of a larger amount of soluble protein and easy removal of the tag at low temperature (Fig. 5.3). More work should also be done on refolding denatured HrpX, which is present in large and highly purified amounts in inclusion bodies.
At this point, we do not know the nature of the ligand(s) that interacts with HrpX. Sequence analysis alone is unable to predict its nature, because of the generally low similarity (around 20-28%) among various
PAS domains and the small number of ligands known so far. Only five crystal or solution structures of PAS proteins are known and the residues important for ligand binding have been identified in only a few of these systems (Repik et al., 2000; Dunham et al., 2003; Gerharz et al., 2003).
The PAS domains most similar to HrpX are found in NodV PAS-A (for
PAS1) and TraJ (for PAS2) and the ligands for these have been described.
NodV senses genistein signals during the interaction with legume plants
(Loh et al., 1997) and its loss leads to restriction of symbiotic host range
(Gottfert et al., 1990). NodV has five tandem PAS domains, but there is no proof that any of them are involved in genistein-sensing. TraJ PAS
290 domains represent an interesting hit. TraJ is a nucleotide-binding outer
membrane protein, unrelated to two-component system kinases, that regulates conjugal transfer of F factors.
In our attempts to identify the growth conditions and signals transduced by HrpX/HrpY, we discovered that HrpY[D57N], when expressed from
multicopy plasmids, could overcome the repression of hrp genes by either
citrate or succinate were supplied as the sole C-source and, to a lesser
extent, by 0.125% tryptone. In contrast, the other environmental
conditions that modulate hrpS expression, such as pH, osmolarity,
nicotinic acid and peptone, were independent of HrpY regulation. The significance of the effect of citrate, succinate and tryptone is still under investigation. Citrate and succinate are important intermediates of the
tricarboxylic acid and glyoxylate cycles. Tryptone, the major component
of LB broth, contains large amounts of free amino acids and small
peptides. Aerobic growth in media containing tryptone as sole carbon and
nitrogen source should activate gluconeogenic and anaplerotic pathways
feeding into the Krebs cycle. Whether the particular amino acid
composition of tryptone leads to a build up of Kreb’s cycle intermediates,
or else, whether the growth in minimal media with succinate or citrate as
sole C-source alter particular amino acid synthesis pathways is not
known. Intriguingly, the periplasmic PAS domain of the E. coli CitA
sensor, which regulates the transport and metabolism of citrate, has
recently been shown to cocrystalize with citric acid molecules bound to
291 the PAS core pocket (Gerharz et al., 2003; Reinelt et al., 2003). By analogy, it is tempting to hypothesize that either citrate or succinate may function as a ligand for one of the HrpX PAS domains, but it is also likely that they may indirectly affect it. In this model the HrpX-citrate conjugate would be locked in a HrpY-phosphatase conformation. At this point, we have only shown modest repression using these two Kreb’s cycle intermediates (only 60 to 40% downregulation compared to the positive control) and it may not be significant from a biological point of view. In our suppression experiments, we tried to control most of the variables, osmotic pressure in particular, by balancing chemical concentrations of ions in the various media tested. Since we could not perform direct osmotic measurements, we used the van t’ Hoff law to approximate their values. When using complex organic substrates such as tryptone, these calculations are impossible. Therefore, in the IM5.5 tryptone treatments some of the repression may be due to the osmotic pressure. This could explain why hrpY[D57N] did not fully overcome the repression (70% suppression) (Fig. 5.12) caused by peptone, because we used 2% peptone versus 0.1% tryptone.
A biochemical approach is probably the best avenue for identifying the molecular ligands for HrpX. If soluble HrpX PAS domains are eventually purified in large amounts, the determination of their solution structure by
292 NMR and the analysis of their chemical shifts in presence of putative cofactors could quickly identify what ligands they bind, as shown for other PAS domains (Amezcua et al., 2002).
From a practical point of view, the study of the biochemistry of HrpX and the identification of its ligands may open the way to large throughput screenings for structurally analogous compounds that could inhibit kinase activation. PAS domains are found in sensors of many other plant and animal pathogens, and, if the molecular basis for their ligand specificity is eventually understood, they could become novel and universal targets for therapeutic drugs and plant disease control chemicals.
5.5 SUMMARY
P. stewartii HrpX/HrpY two-component system activates the hrp/hrc type III secretion system through a regulatory cascade involving the HrpS enhancer-binding protein and the HrpL alternative sigma factor. The HrpX sensor kinase is required for full activation of hrp regulatory and secretion genes in vitro and for consistent infection of corn seedlings. The signal(s) controlling HrpX are not known. The input domain of the sensor is composed of two tandem PAS domains showing little similarity to each others. The presence of PAS domains and the predicted cytoplasmic localization of the sensor suggest that it could be sensing cellular redox levels, the energy charge or other internal metabolic signals.
293 In this study we begun testing the role of the PAS domains in HrpX
activity. We first engineered in frame deletions spanning both individual
motifs and the whole input domain. The effects of the new alleles on
virulence and hrp gene expression were analyzed by corn whorl
inoculations and reporter gene assays. The results suggest that the two
PAS domains are non-redundant, are both required to different extent for
full virulence and a ∆PAS1 deletion had negative effect on virulence when
expressed in a ∆hrpX or ∆(PAS1-PAS2)
We then tried to determine which of the many environmental signals
controlling hrp gene transcriptions are perceived by HrpX. Given that we
were unable to isolate a constitutive HrpX* mutant, we used
overexpression of hrpY[D57N] to attempt to suppress the downregulatory
effects of high pH, high osmolarity, nicotinic acid, complex nitrogen
sources, and some organic acids on a hrpS::lacZ reporter fusion. We
indirectly found that the HrpX/HrpY mediates the effect of tryptone, citrate and succinate but none of the other growth conditions used.
Experiments aimed to establish reagents for the biochemical analysis of
HrpX and the two PAS domains are also presented in this chapter.
294 Strain, or plasmid Relevant phenotype(s) and/or genotype(s) Source or reference
Escherichia coli
DH10B F- mcrA ∆(mrr-hsdRMS-mcrBC) φ80dlacZ∆M15 ∆lacX74 endA1 recA1 deoR
∆(ara,leu)7697 araD139 galU galK nupG rpsL λ- Invitrogen
- - - r HB101 F thi-1 hsd20 (r Bm B) sup E44 recA13 ara-14 leuB6 proA2 lacY1 rpsL20 (Sm )
xyl-5 mtl-1 Boyer and Roulland-Dussoix (1969)
BL21(DE3)/pLysS F- ompT hsdSB (rB- mB-) gal dcm (DE3) carrying pLysS Novagen
- R TB1 F ara ∆(lac-proAB) [φ80 dlac ∆(lacZ)M15] rpsL (Str ) thi hsdR Novagen
- R ER2507 F ara-14 leuB6 fhuA2 ∆(argF-lac)U169 lacY1 glnV44 gal44 galK2 rpsL-20 (Str )
R xyl-5 mtl-5 ∆(malB) zjc::Tn5 (Kan ) ∆(mcrC-mrr)HB101 Novagen
P. stewarti subsp. stewartiii
SS104 Wild type, Wts+, HR+ ICPPBa
DC283 SS104 Nalr Coplin et al. (1986)
DM064 DC283 hrpY1296::Tn5 (Kanr) Frederick et al. (2001)
DM733 DC283 hrpY64::Tn5 hrpS1::Tn3HoHoI (Kanr, Ampr) Merighi et al. (2003)
DM2790 DC283 ∆hrpX17-414 Merighi et al. (2003)
MM254 DC283 hrpY[D57N] Merighi et al. (2003)
MM266 DC283 ∆hrpX15-198 This study
MM284 DC283 ∆hrpX15-270 This study
MM348 DC283 ∆hrpX15-130 This study
MM360 DC283 ∆hrpX143-259 This study
Plasmids pBluescript
KS and SK (+) ColE1 αlacZ (ApR) cloning vector Stratagene pPL6GUSC pLAFR6 derivative carrying a promoterless uidA gene (TcR) Knoop et al. (1991)
R Q pQE30 ColE1 (Ap ) Ptac lacI N-terminal His6-tag expression plasmid Qiagen
pRK415 IncP αlacZ (TcR) Keen et al. (1988)
R Q pTYB1 ColE1 (Ap ), PT7 lacI C-terminal SceVMW intein-CBD tagexpression plasmid NEB
Table 5.1 (continued)
295 Table 5.1 (continued)
r pET15b ColE1 (Ap ) PT7lac lacI N-terminal His6-tag expression plasmid Novagen
r pET28b ColE1 (Km ) PT7lac lacI C-terminal His6-tag expression plasmid Novagen
r pET41b ColE1 PT7lac lacI (Km ) N-terminal GST-S-His-tag expression plasmid Novagen
R Q pEXT20 ColE1 (Ap ) Ptac lacI expression plasmid Wilson et al., 1996
r Q pGEX-2T ColE1 (Ap ) Ptac lacI N-terminal GST- -tag expression plasmid Pharmacia
r Q pMALc2X ColE1 (Ap ) Ptac lacI N-terminal malE fusion expression plasmid NEB
pMM58 pBSK carrying hrpL, hrpXY and hrpS Merighi et al., 2003
+ pMM228 pET15b with a 1.5 kb BamHI hrpX PCR fragment to express a His6-HrpX[1-485] protein fusion This study pMM233 pET15b with a 0.6 kb BamHI ∆hrpX PCR fragment to express His6-HrpX[277-485] protein fusion This study
pMM242 pBSK with a 1.3 kb EcoRI-XbaI fragment carrying a ∆hrpX15-270 allele This study pMM243 pBSK with a 1.3 kb EcoRI-XbaI fragment carrying a ∆hrpX15-198 allele This study
pMM247 pLD55 with a XhoI-SstI insert from pMM243 This study
pMM249 pLD55 with a XhoI-SstI insert from pMM242 This study
pMM251 pET15b with a 0.4 kb BamHI ∆hrpX PCR fragment to express a His6-HrpX[1-142] protein fusion This study pMM257 pET15b with a 0.8 kb BamHI ∆hrpX PCR fragment to express a His6-HrpX[1-276] protein fusion This study pMM265 pET41b with a 0.4 kb BamHI ∆hrpX PCR fragment to express a GST-HrpX[1-142] protein fusion This study pMM270 pET41b with a 0.8 kb BamHI ∆hrpX PCR fragment to express a GST-HrpX[1-276] protein fusion This study pMM280 pET15b with a 0.4 kb BamHI ∆hrpX PCR fragment to express a His6-HrpX[136-276] protein fusion This study pMM283 pBSK with a BamHI-EcoRI insert carrying a SOE-PCR ∆hrpX15-130 allele This study
pMM287 pBSK with a BamHI-EcoRI insert carrying a SOE-PCR ∆hrpX143-259 allele This study
pMM315 pMALc2x with a 1.5 kb BamHI hrpX+ PCR fragment to express a MBP-HrpX[1-485] protein fusion This study
+ pMM319 pEXT20 with the HindIII-XbaI His6-hrpX fragment of pMM228 This study pMM324 pLD55 with a XhoI-SstI insert from pMM283 This study pMM327 pLD55 with a XhoI-SstI insert from pMM287 This study pMM362 pTYB1 with a 1.5 kb BamHI hrpX+ PCR fragment to express a HrpX[1-485]-SceCBD fusion This study pMM363 pTYB1 with a 0.8 kb NheI-XhoI hrpX PCR fragment to express a HrpX[1-276]- SceCBD fusion This study pMM367 pET28b with a 1.5 kb BamHI hrpX+ PCR fragment to express a HrpX[1-485]- His6 protein fusion This study
pMM371 pGEX-2T with a 1.5 kb BamHI hrpX+ PCR fragment to express a GST-HrpX[1-485] protein fusion This study
pMM374 pGEX-2T with a 0.8 kb BamHI hrpX PCR fragment to express a GST-HrpX[1-276] protein fusion This study
pMM377 pQE30 with a 1.5 kb BamHI hrpX+ PCR fragment to express a His6-HrpX[1-485] protein fusion This study pMM378 pRK415 with a BamHI-EcoRI insert from pMM287 This study
Table 5.1 (continued)
296 Table 5.1 (continued) pMM380 pRK415 with a BamHI-EcoRI insert from pMM283 This study
pMM389 pRK415 with a HindIII-EcoRI insert carrying hrpX+ driven by the native promooter This study
Table 5.1 Bacterial strains and plasmids.
297 Sequence coordinates
Primer Sequence (5’Æ 3’) (5’Æ3’)a Source
EdX1 CGCGAATTCTTCGGTATTGCCCTGAACCT 919-938 This study
BdX1 CGCGGATCCTTTGATTGGCGTATCACTGCT 1427-1407 This study
BdX5A CGCGGATCCATGCGCTTGCGTTTGTTAACA 2190-2210 This study
BdX5B CGCGGATCCCATATTGAACGCTTTCTTAAA 974-1994 This study
BdX4AB TCTAGAACGGCCGTTATCACTCACTG 2671-2687 This study
XdX3 CGCTCTAGATTTGCTCGGCGAGCATTTGG 3272-3253 This study
ETXF1 CGCGGATCCGATGCAGATTTTGGACAGAC 1379-1398 This study
ETXF2 CGCGGATCCGACAAGGCAGCGAGAAATATC 2207-2227 This study
ETXR GGATCCTCACTCACTCAGGGTATTAAAGGAATGGTC 2818-2837 This study
ETPAS1R CGCGGATCCCTATCATCTACTGTTCCCGAGCATAAGC 1788-1805 This study
ETPAS2F CGCGGATCCGGGTGTAGCTTATGCTCGGGAACAG 1785-1805 This study
ETPAS2R CGCGGATCCCTATCATCTACAAACGCAAGCGCATAT 2188-2205 This study
XCHISF2 CATGCCATGGAGATGCAGTATTTGGACAGAC 1378-1398 This study
XCHISR2 CCGCTCGAGGGGTATTAAAGGAATGGTC 2816-2834 This study
XBAMF(+1) CGCGGATCCATGCAGTATTTGGACAGAC 1380-1398 This study
XNHEF ACTAGCTAGCATGCAGTATTTGGACAGAC 1380-1398 This study
PAS2XHOR ATGCCTCGAGATGAATGGCTGTGATATC 2142-2159 This study
XXHOIR ATGCCTCGAGGGGTATTAAAGGAATGGTC 2816-2834 This study
dXPASF1 CGGAATTCAGAGGAAGACGGCAGTTATC 1122-1141 This study dXPAS1R1 ACCTGCTCTAGTTGCTTGATTGGCGTATCACTGCTATG 1404-1424 This study dXPAS1F2 AGTGATACGCCAATCAAGCAACTAGAGCAGGTAG 1770-1788 This study dXPASR2 CGGGATCCTCAGTCAGGGTATTAAAGGAATGGTCA 2815-2838 This study
dPAS2R1 CTTCTTTTCAGCGTAATGCTGTTCCCGAGCATAAGCTAC 1805-1785 This study
dPAS2F2 TATGCTCGGGAACAGCATTACGCTGAAAAGAAGTTAG 2157-2178 This study
XPMALR CGGAATTCTCACTCAGGGTATTAAAGGAATGGTC 2816-2837 This study
XPMALF GGGAATTCATGCAGTATTTGGACAGAC 1380-1398 This study
Table 5.2 Oligonucleotide primers.
298
hrpY 2837 hrpX 1380 hrpL
PAS2 PAS1
XdX3 BdX4 BdX1 EdX1 XbaI A BamHI BamHI B EcoRI pDM2842, ∆ pDM2848 1421 919
XdX4AB BdX5B XbaI CDBamHI BamHI EcoRI pMM243, pMM247 ∆ 2690 1974 1421 919
BdX5A XbaI EFBamHI BamHI EcoRI
pMM242, pMM249 ∆
2690 2190
dXPAS1F2 dXPAS1R1 GH BamHI EcoRI pMM283, pMM324 ∆
2837 1770 1425 1122
dXPASR2 dXPAS2F2 dXPAS2R1 dXPASF1 IL BamHI EcoRI pMM287, pMM327 ∆ 2837 2157 1805 1122
PshAI M FspI pMM389, pDM2890, pDM2893 2945 816
HindIII HindIII BamHI
5363 3579 1 pMM58
Figure 5.1. Map of primers and PCR fragments used to construct hrpX mutations
(plasmids and chromosomal mutants).
Plasmids carrying the various alleles are listed on the left and PCR primers are shown by arrowheads. Sequences of the primers are in Table 5.2. Nucleotide coordinates are relative to Genbank accession AF282857
299
DM2790 ∆17−414
MM284 ∆15−270
MM266 ∆15−198
PshAI FspI hrpX H hrpY PAS2 PAS1 hrpL 2837 1380
MM36 MM34 ∆143−259 ∆15−130
PshAI FspI pRK415 Plac pMM389
BamHI EcoRI
pRK415 Plac ∆ pMM378 BamHI EcoRI pRK415 Plac ∆ pMM380
Figure 5.2. Physical maps of the hrpX mutations and plasmids used for
complementation studies.
The coordinates of the deletions refer to the amino acid sequence of HrpX.
300
Figure 5.3. Plasmids used for expression and purification of HrpX.
Lines below the genetic map depict inserts cloned into various expression vectors.
Relevant restriction sites and promoters (P) are indicated. Plasmid names are listed on the right. The nature of the tag used for affinity column chromatography and the plasmid vector name is given in parentheses. Triangles indicate the proteolytic cleavage site for releasing the tag from the recombinant protein. Nucleotide coordinates are relative to Genbank accession AF282857. CBD/Sce = chitin binding/intein domains.
301
2837 hrpX 1380 PAS2 PAS1
ETXR ETXF BamHI BamHI (His tag pET15b) PT7 pMM228 6
ETXR XBAMF(+1) BamHI BamHI
Plac pMM377 (His6 tag pQE30) SG
XcHISR2 XcHISF2 XhoI NcoI
PT7 pMM367 (His6 tag pET28b)
XpMALR XpMALF EcoRI EcoRI
Ptac
Xa pMM315 (MBP tag pMALc2x)
ETXR XBAMF(+1) BamHI BamHI
Ptac
pMM371 (GST tag pGEX-2T)
XNHER XNheF XhoI NheI
CBD Sce PT7
SSGEL pMM362 (Intein-CB tag pTYB1)
302
Figure 5.4. Plasmids used for expression and purification of HrpX domains.
Lines below the genetic map depict inserts cloned into various expression vectors.
Relevant restriction sites and promoters (P) are indicated. Plasmid names are listed on the right. The nature of the tag used for affinity column chromatography and the plasmid vector name is given in parentheses. Triangles indicate the proteolytic cleavage site for releasing the tag from the recombinant protein. Nucleotide coordinates are relative to Genbank accession AF282857. CBD/Sce = chitin binding/intein domains.
303 2837 hrpX 1380 PAS2 PAS1
BamHI BamHI
PT7 pMM257 (His6 tag pET15b)
BamHI BamHI
PT7 pMM251 (His6 tag pET15b) BamHI BamHI
PT7 pMM280 (His6 tag pET15b) BamHI BamHI
PT7 pMM233 (His6 tag pET15b)
BamHI BamHI
Ptac
pMM270 (GST tag pET41b)
BamHI BamHI
Ptac
pMM265 (GST tag pET41b)
XhoI NheI
CBD Sce PT7 pMM363 (Intein-CB tag pTYB1) SSGEL MSAM
BamHI BamHI
Ptac SGR pMM374 (GST tag pGEX-2T)
304
P. agglomerans pv. gypsophilae P. stewartii E. amylovora
P. chrysanthemi
P. syringae*
P. aeruginosa* X. campestris
0.1
Figure 5.5. Phylogenetic tree of HrpX homologs.
HrpX orthologs, or presumed orthologs (*) found in the genome of P. stewartii, P.
agglomerans pv. gypsophiliae, P. chrysanthemum, E. amylovora, X. campestris, P.
syringae (PSPTO559; gi28851022) and P. aeruginosa (PAO600; gi9946473) were
aligned with ClustalX. Other accession numbers are shown in Table 2.3. Large gaps,
if any, were eliminated and a phylogenetic tree was constructed with the neighbor-
joining methods and 1000 bootstraps cycles. The unrooted tree is shown. Each branch
was supported by a bootstrap value >980. The bar indicates a genetic distance of 0.1.
305
Figure 5.6. PSI-PRED Secondary structure analysis of HrpX.
The amino acid sequence of HrpX was analyzed with PSI_PRED at http://bioinf.cs.ucl.ac.uk/ to identify the secondary structure elements. PAS domains are labeled following the convention of Amezcua et al., (2002). Conserved motifs in the transmitter domain were identified according to Hoch et al., (1995) and Stock et al., (1989). α-helices are shown as green cylinders, β-sheets are represented by yellow arrows and loops are shown as black lines.
306
PAS1
Aβ Bβ
Cα Dα Eα Fα
Gβ Hβ Iβ
PAS2
Aβ Bβ
Cα D/Eα? Fα
Gβ Hβ
linker
(continued)
307 Figure 5.6. (continued)
H~P H box
H-ATPase domain
N box D box
G box
α-helix CONF: CONFIDENCE OF PREDICTION β-strand Pred: prediction of secondary structure coil
Fig.B. PSI-PRED Secondary structure analysis of HrpX
308
PHYLIP_1 consensus
128494 NodV PAS1 PAS1
gi|2494692 WC1 gi|3041695 KinA gi|120207 FixL
6980436
8569635 gi|2145515 FixL
128594 NtrB gi|128596 NtrB 1171792 NtrB
gi|729901 KinC gi|1168485 ArcB gi|585587 NtrY
5107751 0.1
Figure 5.7. Phylogenetic tree of PAS1-like domains in other two component systems
The 14 PAS domains most similar to the P. stewartii PAS1 core domain were identified by a CDD search at NCBI and aligned with ClustalX. Accession numbers are shown next to the protein names. Large gaps were eliminated and a phylogenetic tree was constructed with the neighbor-joining method and 1000 bootstrap cycles.
TreeView was used to construct the cladogram above. The bar indicates a genetic distance of 0.1.
309
10 20 30 ------CCCEEEEEECCCCEEEECHHHHHHHCCCHHHHCCCC 1drmA0 ------IPDAMIVIDGHGIIQLFSTAAERLFGWSELEAIGQN * || || | |* |*| | * HrpX MQYLDRRLHSSDTPIKLLEFAMNMLRDAVYLIDSDTCFFYVNDEACRMLGYSNAELLNMR CHHHHHHHHHHHHHHHHHHHHHHHCCCCEEEECCCCCEEECCHHHHHHHCCCHHHHHCCC 10 20 30 40 50 60
40 50 60 70 80 90 HHHHCCCCHHHHHHHHHHHHHHHCCCCCCCCCEEEEEECCCCCEEEEEEEEEEEEECCEE 1drmA0 VNILMPEPDRSRHDSYISRYRTTSDPHIIGIGRIVTGKRRDGTTFPMHLSIGEMQSGGEP | * || | | | | | *| HrpX VSDIDPEWDFDDVMDMWKRVREQNGGR--PFTFETFHKSSQGEMIAVEISANPFVYEEKS HHHCCCCCCHHHHHHHHHHHHHHCCCC--EEEEEEEEECCCCCEEEEEEEEECCCCCCEE 70 80 90 100 110
100 110 EEEEEEEECHHHHHHHHHHHHCC------1drmA0 YFTGFVRDLTEHQQTQARLQELQ------| |*|* | | * * HrpX YSMCVVKDIRERKQLEQVAYAREQEFRVLVENSPDMVVRFSPDLKCQYANPAALRHLRLS EEEEEECCCCCCEEEEEEECCCCCCEEEEECCCCEEEEEECCCCEEEECCHHHHHHHCCC 130 140 150 160 170
Figure 5.8. Threading of the PAS1 amino acid sequence into the FixL PAS secondary structure.
The PAS1 domain of HrpX was threaded into the B. japonicum FixL PAS domain
using the mGenTREADER algorithm at http://bioinf.cs.ucl.ac.uk/. α-helical regions
are indicated by the letter H. β-sheets are labeled with the letter E. Random coil loops
are indicated by Cs. Identical residues are indicated with (|) and similar residues are
marked by (*).
310
Helical connector (Fα)
PAS core
β-scaffold
Ligand binding pocket
E67 P66 G100 K96 R81 V61
R79 E55 Eα Fα S52 Dα
Gβ G50 Bβ Cα V104 D33 Aβ Eβ Hβ
Y97 V102 I32 D29 A45 R47 S108 D28
Figure 5.9. Structural model of the P. stewartii PAS1 threaded into the backbone of B. japonicum FixL PAS.
311
Figure 5.10. Complementation analysis and virulence phenotypes of P. stewartii hrpX deletion mutants.
Inocula were prepared and plants were inoculated as described in Material and
Methods. Bacterial suspensions were adjusted to A600nm = 0.1. Whorls of 6-day-old corn seedlings were inoculated and watersoaking symptoms were scored at 4 days using a 0 to 3 scale. The cartoons on the left illustrate which part of the HrpX protein was deleted in the various mutants. Strains: DC283 is wild-type; DM2790 is
∆hrpX17−414, MM266 is ∆hrpX 15−198, MM284 is ∆hrpX 15−270, MM348 is ∆hrpX
15−130, MM360 is ∆hrpX 143−259. Plasmids: in all plasmids the expression of the gene is driven by the native hrpX promoter; pMM389 carries hrpX+ , pMM380 carries a ∆hrpX
15−130 allele and pMM378 carries ∆hrpX 143−259.
312 .
1 2 ~ DC283 MM360/pMM378 MM360/pMM380 1 ∆ ~ MM360/pMM389 MM360 MM348/pMM378 MM348/pMM380 ∆ 2 ~ MM348/pMM389 MM348
MM284/pMM378 A MM284/pMM380 ∆ ~ MM284/pMM389 MM284 MM266/pMM378 MM266/pMM380 ∆ ~ MM266/pMM389 MM266 DM2790/pMM378 DM2790/pMM380 ∆~ DM2790/pMM389 DM2790
00.511.522.53 Watersoaking
313
100 90 80 70 60 GUS activity % 50 Wts%
Percent 40 30 20 10 0
/2531 M2531 M2531 M2531 254 66/pD 48/pD 2 3 MM DC283/pDM2531M2790/pD D MM MM284/pDM2531MM MM360/pDM2531
Figure 5.11. The effect of various hrpX mutations on virulence and the expression of hrpJ. a typical hrp secretion gene.
Expression of the hrpJ-lacZ reporter gene fusion in pDM2531 was measured in P.
stewartii strains resuspended in IM5.5 to an A600=0.05-0.1 and grown for 16 h at
28°C. Data are from two sets of experiments with at least two replicates per strain.
Error bars indicate standard deviations (when not visible is because the SD is too
small for the scale of the graph). All data are given as a percentage of wild-type
levels. DM2790 is ∆hrpX17−414, MM266 is ∆hrpX15−198, MM284 is ∆hrpX15−270, MM348 is
∆hrpX15−130, MM360 is ∆hrpX143−259, MM254 is hrpY[D57N]. MM254(pDM2531) had
no GUS activity or virulence
314
Figure 5.12. The effect of environmental stimuli on hrpS-lacZ expression and suppression of these effects by ectopic overexpression of hrpY[D57N]
P. stewartii strains MEX1 hrpS-lacZ and its derivative DM733 hrpY::Tn5 hrpS-lacZ carrying pMM118 Plac-hrpY[D57N] were resuspended in IM5.5+supplement (initial
A 600nm = ~0.2-0.3) and grown at 28C for 12-16 h. Expression of hrpS-lacZ was measured and the data are shown as a percentage of the respective control strain grown in IM. Media: IM5.5A was standard IM inducing media lacking casaminoacids and with 10 mM sucrose as the C-source; “high Pi ” was IM5.5A with 250 mM
KH2 PO4 ; “High NaCl” was IM5.5A with 250 mM NaCl; “pH8.0” was IM buffered with
HEPES to pH 8.0; “citrate” was IM5.5A with 10 mM trisodium citrate as a sole C-
source; “succinate” was IM5.5A with 13 mM succinic acid as a sole C-source;
“asparagine” was IM5.5A with 56 mM L–asparagine as sole source for C and N;
“peptone” was IM5.5A with 2.0% peptone as a sole source for C and N; “tryptone”
was IM5.5A with 0.125% tryptone as sole source of C and N; “tryptone-sucrose” was
IM5.5A with 0.125% tryptone as sole N source. The pH of the media was adjusted
with NaOH and the milliequivalents of base were included in calculations of osmotic
pressures. Error bar = SD.
315 120 IM5.5 High Pi 100 High NaCl pH8.0 80 citrate succinate 60 asparagine
% IM5.5 control peptone 40 tryptone tryptone-sucrose 20 LB Nicotinic acid 0 MEX1 DM733/pMM118
316
Figure 5.13. Pilot experiment to overexpress polyHis-tagged HrpX proteins
E. coli BL21(DE3) pLysS cells carrying various expression plasmids were grown in
LB to A600=0.6 and induced with 1 mM IPTG for 3 h. Cells were concentrated 10- fold in PBS and lysed with reducing sample buffer. Total cell proteins (TCP) were run in a Laemmli gel. Lane 1, prestained molecular weight marker (Invitrogen), l, lane 2-
3, uninduced-IPTG induced cells carrying pMM223 (His6 -HrpS); lane 4-5, uninduced-
IPTG induced cells carrying pMM228 (His6 -HrpX); lane 6-7, uninduced-IPTG induced
cells carrying pMM233 (His6 -HKHrpX); lane 8-9, uninduced-IPTG induced cells
carrying pMM251 (His6 -PAS1); lane 10-11, uninduced-IPTG induced cells carrying pMM257 (His6 -PAS1-2). Subsequent analysis has shown that in all cases the proteins
precipitate in inclusion bodies.
317
1 2 3 4 5 6 7 8 9 10 11
78 kDa His -HrpX (lane 5) 61 kDa 6
His6-HrpS (lane 3)
His6-PAS1-2 (lane 11)
His6-HK (lane 11) 25 kDa
His6-PAS1 (lane 9) 13 kDa
318
Figure 5.14. Overexpression and purification of His6 -HrpX from E. coli
BL21(DE3) (pMM228).
(A) Coomassie-stained gel showing various steps in the purification of His6 -HrpX:
Lane 1, molecular weight markers (Invitrogen); lane 2, total cell protein (TCP) from uninduced cells; lane 3, TCP from IPTG-induced cells; lanes 4 to 10, eluted fractions at 40 mM, 60mM, 80 mM, 100 mM, 150 mM, 250 mM, and 500 mM imidazole. (B)
Anti-poly(His)6 Western blot analysis of purified fractions: lane 1, TCP from lane 2, panel A; lanes 2 to 4, fractions eluted at 40 mM, 100 mM and 150 mM imidazole. (C)
Fractions from a separate purification (used for MS/MS protein fingerprinting) eluted at 500 mM imidazole after extensive washing in 80 mM imidazole: Lane 1, molecular weight markers; lane 2, HrpX fraction in 500 mM imidazole. (D) Anti-poly(His)6
Western blot analysis of fractions from the purification of the HrpX preparation used for MS/MS analysis: Lane 1, TCP from uninduced cells (10X); lane 2, TCP from
IPTG-induced cells (10X); lane 3, cleared lysate from IPTG-induced cells loaded onto the Ni2+-NTA columns (30X); lane 4, inclusion bodies (30X); lane 5, column flow- through; lane 6, 500 mM imidazole eluted fraction (same as lane 2, panel C).
319 A 1 2 3 4 5 6 7 8 9 10
61 kDa His6-HrpX
B
α-Poly(His)6 1 2 3 4
His6-HrpX 61 kDa
C D 1 2
1 2 3 4 5 6 7 α-Poly(His)6
61 kDa
36 kDa 19 kDa
320
Figure 5.15. Example of the results from nano LC-MS/MS protein fingerprinting of the tryptic fragment LHSSDTPIK from gel-excised His6 -HrpX.
321
Figure 5.16. Overexpression and purification of MBP-HrpX by amylose affinity and ion exchange chromatography.
(A) Purification by amylose affinity chromatography from strain DH10B (pMM315):
Lane 1, prestained molecular weight marker (Invitrogen); lane 2, TCP from uninduced cells; lane 3, TCP from IPTG-induced cells; lane 4, amylose columns flow-through; lanes 5-6, wash fractions; lanes 7-15, eluted fractions using 10 mM maltose. (B)
Factor Xa proteolysis: Lane 1, prestained molecular weight marker (Invitrogen); lanes
2-4, various 10 mM maltose fractions from Panel A after pooling, ultrafiltration and storage at 4°C for 16 h; lane 5, undigested MBP-HrpX preparation (note again the presence of degradation fragments); lane 6, MBP-HrpX preparation from lane 5 digested for 6 h at 22°C with factor Xa to release the 56 kDa HrpX protein. (C) Ion exchange chromatography: lane 1, prestained molecular weight marker (Invitrogen); lanes 2-13, protein preparations digested with factor Xa for 24 h and purified by ion- exchange chromatography in a Q-sepharose column using a linear gradient of NaCl from 20 mM to 750 mM. (fractions from 20 to 320 mM are shown in the gel). (D).
Summary of MBP-HrpX purification steps: Lane 1, prestained molecular weight marker (Invitrogen); lane 2, TCP from uninduced cells; lane 3, TCP from IPTG- induced cells; lane 4, soluble total proteins; lane 5, flow-though from the amylose column; lane 6, concentrated 10 mM amylose eluted fraction; lane 7, Factor Xa digested preparation; lane 8, anion echange chromatography purified concentrated fraction.
322 A 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 113 kDa
81 kDa
1 2 3 4 5 6 7 B
63 kDa 56 kDa
C 1 2 3 4 5 6 7 8 9 10 11 12 13
63 kDa
D 1 2 3 4 5 6 7 8
63 kDa 56 kDa
323
A 0 2 5 10 15 30 60 min
HrpX~32P
B 160000
140000
120000
100000
80000
60000 Sum of pixels
40000
20000
0 0 20406080 minutes
32 Figure 5.17. Kinetics of autophosphorylation of His6 -HrpX by [γ P]ATP.
(A) Reaction mixtures containing 30 µM His6 -HrpX were incubated with 0.22 µM µCi
[γ 32P]ATP for increasing times as indicated in the figure. Reactions were terminated with stop sample buffer and separated in a Laemmli gel for subsequent autoradiography of the radiolabeled proteins. (B) The radioactive signals were quantified after exposure to a Phosphor storage cassette. The signals are reported in the plot as arbitrary units (sum of pixels in a defined constant sampling area).
324
Incubation time: 60 60 0 2 5 10 15 30 60 60 min
1 2 3 4 5 6 7 8 9 10 A 89 kDa
26.1 kDa
B 89 kDa 58 kDa BarA~32P HrpX~32P Background kinase
26.1 kDa HrpY~32P
32 32 Figure 5.18. Transphosphorylation of HrpY by BarA~ P and His6 -HrpX~ P.
Coomassie-stained PAGE gel (A) and autoradiography (B) of reaction mixtures containing [γ 32P]ATP and the following semipurified proteins: lane 1, BarA; lane 2,
25 pmol His6 -HrpX; lane 3, 380 pmol His6-HrpY; lanes 4 to 9, His6 -HrpX and His6 -
HrpY (1:15 molar ratio) incubated for 0, 2, 5, 10, 30 and 60 min; lane 10, BarA and
His6 -HrpY (1:1 molar ratio). Note the presence of a copurifying ~35-kDa uncharacterized background kinase in all preparations. The His6 -HrpY preparation also showed a moderate intensity signal at 26.1 kDa in the absence of any kinase. This observation prompted further purification of the His6 -HrpY protein by ion-exchange
chromatography as described in Chapter 4. The higher intensity signal at 26.1 kDa in
lanes 6 and 7 suggest a low level of phosphorylation of HrpY by HrpX~P.
325 A 1 2 3 4
66 kDa
B 1 2 3 4
Background kinase 66 kDa ? Background kinase
Figure 5.19. Purification of the GST-PAS1-2 protein from E. coli DH10B
(pMM374) and attempted direct photoaffinity labeling to [γ 32P]ATP.
(A) Overexpression and purification of the GST-PAS1-2 peptide containing both HrpX
PAS domains by glutathione affinity chromatography: lane 1, TCP from uninduced cells; lane 2, TCP from IPTG-induced cells, lane 3, cleared lysate; lane 4, fraction eluted from the GST-BindTM column with 10 mM reduced glutathione. (B) Direct
photoaffinity labeling of GST-PAS1-2 with [γ 32P]ATP. Lanes 1 to 4, GST-PAS1-2
preparation shown in Panel A, lane 4 incubated with [γ 32P]ATP under UV light for 5,
15, 30 and 45 min, respectively. The BSA negative control produced a smear (not
shown). A GST control was not available for this experiment. Note the presence of
background kinases not visible in the Coumassie gel of panel A, lane 4. The arrow
with a question mark indicates a possible PAS1-2~P signal, but the presence of
background kinases will require further purification to confirm it.
326
CHAPTER 6
N-ACYLHOMOSERINE LACTONE-DEPENDENT QUORUM-
SENSING IS REQUIRED FOR FULL EXPRESSION OF THE HRP-
TYPE III SECRETION SYSTEM OF PANTOEA STEWARTII SUBSP.
STEWARTII
6.1 INTRODUCTION
Quorum sensing is the phenomenon by which bacterial populations
coordinate the expression of complex phenotypes in response to their population density (Fuqua et al., 1994). In more than 30 gram-negative
species, quorum sensing is mediated by N-acyl-homoserine lactone (AHL)
signals that are perceived by receptor proteins acting as transcriptional
factors. AHLs are constitutively synthesized and can usually move freely
across biological membranes. The AHL concentration outside of the cell
is proportional to the population density in the milieu. When critical
concentrations are reached (>>Kd at equilibrium), AHL binds
quantitatively to the receptor/transcription factor thereby modulating its
affinity for DNA regulatory sites. Many review articles are available on
this subject (Fuqua et al., 1996; Fuqua and Greenberg, 1998; Dunny and 327 Winans, 1999; Parsek and Greenberg, 2000; Bassler and Schauder, 2001;
Miller and Bassler, 2001; Loh et al., 2002; Von Bodman et al., 2003).
AHL-independent quorum sensing mechanisms in gram-negative bacteria have also been described (Surette et al., 1999; Schauder et al., 2001).
Among the bacterial functions regulated by AHLs are: biofilm formation and light production in Vibrio spp. (Nealson et al., 1970; Camara et al.,
2002), toxins and other virulence factors in Pseudomonas aeruginosa
(Passador et al., 1993; Wagner et al., 2003), conjugal transfer of virulence plasmids in Agrobacterium tumefaciens (Piper et al., 1993), secondary metabolites and antibiotic production in Pseudomonas aureofaciens and Pectobacterium carotovorum (McGowan et al., 1995;
Pierson et al., 1998), and exoenzymes and harpinEcc in Pectobacterium
carotovorum (Pirhonen et al., 1993; Cui et al., 1996; Chatterjee et al.,
2002). Often, more than one quorum sensing system operates in the same cell (Bassler and Schauder, 2001). These regulatory networks may be organized into parallel-converging (Mok et al., 2003), parallel- independent (Zhang and Pierson, 2001) or hierarchical pathways, such those described in P. aeruginosa (Pearson et al., 1995). Quorum sensing systems interact with various housekeeping global regulators, such as
RpoN (Lilley and Bassler, 2000), RpoS (Whiteley et al., 2000) and rsmA/rsmB (=csrA/csrB) (Cui et al., 1995; Koiv and Mae, 2001; Pessi et al., 2001), or with two-component systems (Sperandio et al., 2002;
Ledgham et al., 2003; Ledgham et al., 2003). 328 In the plant macerogenic bacterium P. carotovorum subsp. carotovorum,
a complex regulatory network, involving quorum sensing, coregulates Pel,
Peh, Cel and Prt exoenzyme production, carbapenem antibiotic synthesis and the hrp genes.
In Pantoea stewartii subsp. stewartii, which causes Stewart’s wilt and leaf blight of sweet corn and maize, pathogenicity depends on a type III secretion (TTS) system encoded by hrp genes (Alfano and Collmer, 1997).
The Hrp phenotype in this species is defined as the lack of virulence in corn and the inability to elicit an HR upon infiltration of high doses of
IM-induced cells into tobacco leaves (Frederick et al., 2001). In addition, extracellular polysaccharide (EPS) is also a primary virulence factor, being required for wilt symptoms, but not for water-soaked lesion formation on young leaves (reviewed in Leigh and Coplin, 1992). The
EsaI/R quorum sensing system of P. stewartii controls synthesis of EPS
(von Bodman and Farrand, 1995; reviewed in Von Bodman et al., 2003).
The esaI gene encoding the synthase for the production of AHLs and the esaR gene that encodes its cognate AHL-receptor/transcription regulator have been well characterized. Mutations in esaI have pleiotropic effects, including elimination of AHL production, non-mucoidy, reduced growth rates and decreased virulence (von Bodman and Farrand, 1995). The major
AHL produced by P. stewartii is N-3-oxohexanoyl-L-homoserine lactone
(OHHL); subnanomolar concentrations of N-3-oxooctanoyl-L-HSL are also made (von Bodman and Farrand, 1995; Von Bodman et al., 1998). In 329 contrast to the paradigm for LuxR-like receptors (Fuqua et al., 1994),
EsaR negatively regulates EPS production at low population-density (Von
Bodman et al., 1998 by acting as a repressor in the apoprotein
conformation. Binding of the cognate AHL relieves the repression by
decreasing the affinity of EsaR for its DNA binding sites at the esaR promoter (Minogue et al., 2002), the only target DNA site for EsaR identified so far. On the other hand, the regulatory cascade leading to the esaR/esaI-dependent activation/derepression of the EPS-producing cps gene cluster is not known, and experiments in S. von Bodman laboratory have not shown in vitro binding of semipurified EsaR to cps promoter fragments. A putative EsaR-binding site in front of esaR
(GCCTGTCTA:TAGTGCAGGT) partially resembles the luxR-box
C G T T T consensus ( / ANCT / ANCNNNTAN / NG / NCAG / NG) (S. von Bodman,
pers. commun.).
The virulence of the esaI mutant was originally tested using a stem-
inoculation technique. Under these conditions, the esaI strain was unable
to cause the typical water-soaked lesions and wilt observed with wild-type
strains (Von Bodman et al., 1998). Lack of EPS usually leads only to loss
of the ability to move systemically in the plant and cause severe wilting;
most non-mucoid cps mutants can still produce water-soaked lesions on
the pseudostem (Dolph et al., 1988; Von Bodman et al., 1998). More
relevant assays such whorl inoculation (which test specifically the ability
to elicit watersoaking) had not been done with the esaI strain at the start 330 of this study. The apparent lack of watersoaking prompted us to examine
the effect of the Esa quorum sensing system on the expression of hrp
genes.
As shown in previous work (Coplin et al., 1986; Coplin et al., 1992;
Coplin et al., 1992; Frederick et al., 2001) and in previous chapters of
this dissertation, P. stewartii needs to fully express the Hrp TTS system
and effector genes to cause watersoaking in corn.
We therefore hypothesized that hrp expression was repressed in the
esaI mutant and that the regulatory effect could be mediated by the
cognate EsaR receptor. Given that the regulatory Hrp cascade is extremely
complex (Frederick et al., 1993; Merighi et al., 2003), quorum sensing
regulation could enter the pathway at any of many levels: hrpXY, hrpS or
hrpL transcription, HrpX/HrpY post-translational activation, or secretion
and effector gene transcription.
The results of this study showed that P. stewartii esaI mutants have a
Hrp- phenotype at low inoculum doses and that this mutation significantly reduces expression of hrp genes. We also found that transcriptional regulation of hrpS is the first step in the hrp regulatory cascade that is
affected by the esaI mutation. Our findings are consistent with the
hypothesis that EsaR is mediating the effects of esaI. The presence of a
putative esaR box in the 5’-regulatory region of hrpS may indicate direct
regulation. The relevance of quorum sensing to the biology of P. stewartii
will be discussed. 331
6.2 MATERIALS AND METHODS
6.2.1 Bacterial strains and plasmids
The Escherichia coli and P. stewartii strains and plasmids used in this study are listed in Table 6.1. Luria-Bertani (LB) broth and agar (Ausubel
et al. 1987) were used for strain maintenance and for growth of P.
stewartii under Hrp-repressing conditions. To induce hrp genes in P.
stewartii strains, overnight LB broth cultures were washed twice in pH
5.5 inducing medium (IM5.5; Frederick et al. 2001), inoculated into 2 ml
of the same medium at A600nm = 0.01 and grown for 12-16 h, unless
otherwise specified. The final optical density ranged from A600nm = 0.6 to
0.9. For time-course experiments, bacteria were grown in 250-ml flasks with side arms and optical density readings were taken with a Spectronic
20 spectrophotometer (Thermo Spectronic). Liquid cultures were grown in flasks or tubes shaken at 200 rpm at 37°C for E. coli or 28°C for P.
stewartii. When appropriate, antibiotics were supplied at the following
concentrations: ampicillin, 200 µg ml-1; kanamycin, 50 µg ml-1; and tetracycline, 10 µg ml-1. Synthetic N-3-oxo-hexanoyl-homoserine lactone
(OHHL; Sigma) was resuspended in methanol at 13 mM and stored at –20
°C. OHHL was used at a working concentration of 4 or 10 µM.
332
6.2.2 Genetic and molecular biology techniques
DNA isolation, agarose gel electrophoresis, restriction enzyme
digestion and ligation were done according to standard procedures
(Ausubel et al. 1987). PCRs used either native Taq (for colony-PCR
screens and construct mapping) or cloned Pfu (for molecular cloning)
DNA polymerases (Invitrogen and Clontech respectively) and standard amplification protocols (Ausubel et al., 1987). For colony-PCR, we used
cells lysed in 50 mM NaOH for 10 min at 100°C as templates. Plasmids
were introduced into E. coli and P. stewartii strains by electroporation or
mobilized by triparental mating using pRK2013::Tn7 as a helper plasmid
(Ditta et al. 1980).
6.2.3 Plasmid construction
For complementation and over-expression experiments, the hrpY and
hrpL ORFs, including their ribosomal binding sites, were amplified by
PCR and cloned into high and low copy number cloning vectors, such that
the inserts were expressed from the Plac promoter in the vector (Fig. 6.1;
the construction of these plasmids is described in Chapter 2).
Construction of pPL6GUSC plasmids containing uidA reporter genes fused
to the hrpX, hrpS, hrpL, hrpJ and wtsE promoter regions is described in
Chapter 2.
333 6.2.4 Enzyme assays
β-glucuronidase (GUS) activity was assayed fluorometrically using 4-
methyl-umbelliferyl-β-D-glucuronide as described by Jefferson (1987).
However, the assays were scaled down to fit microtiter plates and
analyzed using a Victor 14202 multilabel reader (PE-Applied Biosystems,
Wellesley, MA, U.S.A.) as described in Chapter 2. Net GUS activity of
each strain was corrected for the basal fluorescence of P. stewartii DC283
carrying pL6GUSC without an insert. Specific activity was expressed in
-1 -1 -1 GUS units (pmol MU min OD600nm ml of culture at 37°C).
6.2.5 Plant assays
To test for HR elicitation in tobacco (Nicotiana tabacum L., cv. “White
Burley”) plants, P. stewartii strains were grown overnight in IM5.5 liquid
9 medium, washed and resuspended in water at OD600nm = 0.52 (ca. 1x10
CFU ml-1). Cells were infiltrated into tobacco leaves using two replicates
per strain as previously described (Frederick et al. 2001) and necrosis was
rated 24-48 h after infiltration. Pathogenicity tests were performed by
inoculating the whorls of 6-day-old sweet corn seedlings (Zea mays var.
saccharata, cv. “Seneca Horizon”) as previously described (Coplin et al.
1986). This assay tests specifically for water-soaking ability. The plants
were held in growth chambers at 29°C (photoperiod 16 h, 15,000 lux,
relative humidity 99%). After 3 days, disease severity was rated using a 0
334 to 3 scale (0= no symptoms, 1=scattered small lesions, 2 = numerous
lesions, and 3 = extensive lesions that remained water-soaked with ooze
forming on leaf surfaces).
6.3 RESULTS
6.3.1 An esaI mutant, containing a mutation in the AHL synthase gene,
shows a Hrp- phenotype.
The esaI mutant used in this study, ESN51, is defective in N-3-oxo-C6-
HSL synthesis (OHHL) and EPS production and is unable to cause wilting
of corn (von Bodman and Farrand, 1995). We were initially interested in confirming whether this mutant was indeed unable to produce water- soaked lesions as previous stem-inoculations had suggested (Von Bodman
et al., 1998). To this purpose, we performed repeated corn whorl
inoculations using low inoculum doses (about 5x107 CFU per whorl) of various quorum sensing mutants. The ES∆R ∆esaR::cat and ES∆IR
∆esaIR::cat mutants produced water-soaked lesions that were basically undistinguishable from the wild-type strain after 3 to 4 days-post- inoculation (DPI) (Table 6.2). ESN51, the esaI strain, produced from none to a few scattered small lesions (Table 6.2) and some sporadic leaf flecking. To test if the esa mutations affected the ability to elicit a non- host HR, the mutants were also infiltrated into tobacco leaves. The ability to cause an HR was positively correlated with virulence (Table 6.2), confirming the Hrp phenotype of the esaI strain. 335
6.3.2 In epistasis experiments, constitutive expression of hrpS, but not hrpY, suppresses the Hrp phenotype of esaI strains
To verify the epistatic interaction between hrp and esa genes, we tested the ability of several known hrp regulatory genes to restore virulence to the esaI mutant. Clones containing hrpY, hrpS and hrpL expressed from a constitutive promoter (Plac) (Fig.6.1) were introduced into strains ESN51,
ES∆R and ES∆IR. As shown in Table 6.2, ectopic expression of hrpS and hrpL, but not hrpY, suppressed the Hrp- phenotype of the esaI strain.
Considering that one known role of HrpY is to activate the transcriptional expression of hrpS (Merighi et al., 2003), these results suggest that esaI is epistatic to hrpY. In other words, this implies the existence of regulatory interactions between the hrp genes and quorum sensing.
6.3.3 Expression of most hrp/wts genes is dramatically reduced in the
N-acyl-homoserine synthase mutant
To determine if reduced expression of hrp genes is responsible for the
Hrp- phenotype of the esaI strain and to identify which hrp promoter responds to quorum sensing signals, several plasmid borne hrp-uidA fusions were introduced into strain ESN51 and GUS activity was measured after growth in IM5.5 to late exponential/early stationary phase
(Fig. 6.2). Fusions carried by pMM50 hrpS-uidA, pDM2785 hrpL-uidA, pDM2531 hrpJ-uidA and pDM2791 wtsE-uidA were all downregulated 336 from 10 to 200 fold. However, the hrpX-uidA fusion (pMM25) exhibited only a two-fold difference. The modest effect of quorum sensing on PhrpX may be due to the decreased autoregulation of hrpXY by HrpY, which would in turn be a consequence of the autoregulatory loops described in
Chapter 3. Given this assumption, our results indicate that quorum sensing-dependent regulation of the hrp regulon probably enters the cascade at the level of hrpS regulation, since it is the earliest gene in the pathway to be significantly affected (about 12-fold).
To verify that the effect of esaI mutations on hrp gene expression was due to the defect in OHHL production and not to second-site mutations, we added synthetic OHHL, at a concentration comparable to that observed previously in P. stewartii cultures by Von Bodman et al., 1998, to ESN51 carrying the hrpJ-uidA reporter. hrpJ expression was restored to wild- type levels indicating that the esaI mutation can be complemented by synthetic OHHL (Fig. 6.3).
6.3.4 Expression of hrp secretion genes is population density- dependent and EsaR mediates the effects of esaI
Quorum-sensing regulated genes are expected to differ widely in their expression at various cell densities. This may complicate comparisons of expression data between samples with different cell density. We decided to more carefully analyze the population density dependence of hrpJ, which has been used throughout this dissertation as a representative 337 secretion gene. This was done in wild-type, esaI, esaR and esaIR strains
(Fig. 6.4). Cells were first grown in LB and then diluted 1:200 into fresh
medium. The GUS activity of the hrpJ-uidA reporter (in plasmid
pDM2531) was measured at different cell densities and data were
normalized per A600nm. (The relationship between A600nm and viable cells was the same for all strains used.) As shown in Fig. 6.4, in the wild-type background (strain DC283) the response curve follows a logistic trend, with a quorum for hrp expression estimated around 1 x 108 CFU ml-1 (A quorum was defined as the population density where the second derivative of the logistic curve equal zero; not shown) On the other hand, hrp gene expression in ESN51 was suppressed and basically constant at different cell densities, with only a slight rise at very high populations (Fig. 6.4).
This confirmed that esaI is absolutely required for full hrp gene expression over a wide range of cell densities. Addition of synthetic
OHHL (Sigma) at 10 µM to the growth medium did not up-regulate hrp gene expression in the wild-type strain at low cell densities (strain
DC283(pDM2531), Fig 6.5), which would be expected if quorum sensing was the only mechanism controlling hrp genes. This is in contrast to what was previously observed for EPS production, which was upregulated by exogenous AHL at low cell densities (von Bodman and Farrand, 1995).
EsaR is the only known cognate receptor protein for OHHL in P. stewartii. To test whether esaR mediates the regulatory effect of esaI on hrp genes, we evaluated the expression of a plasmid-borne hrpJ-uidA 338 fusion in a ∆esaRI::cat mutant. In this double esa mutant, hrp gene
expression was restored to levels comparable to wild-type or ∆esaR::cat
strains. For reason we do not fully understand, the ∆esaRI::cat strain showed somewhat lower hrp transcription levels than the ∆esaR::cat mutant, which was mildly upregulated at higher cell densities compared to the wild-type (Fig. 6.4). Collectively, these and the previous results suggest that esaR is either directly or indirectly involved in the repression of hrp genes and that additional regulators may be required. These additional regulators must be expressed only at later growth stages either downstream of or, more likely, in parallel with EsaR. Supporting this model, expression of hrpJ was not upregulated at low cell density in the
∆esaR::cat mutant (Fig. 6.4).
6.4 DISCUSSION
The results of this study show that the esaI/esaR quorum sensing system plays an important role in modulating P. stewartii virulence, irrespective of its effects on EPS synthesis. In this study we showed that a deletion mutation in the OHHL-synthase gene caused a significant reduction in water-soaked lesion formation, HR elicitation and in vitro hrp gene expression. The Hrp phenotype and the regulatory effects of the esaI mutation were suppressed by a second mutation in esaR, suggesting the involvement of the EsaR receptor as a repressor of hrp genes (Fig. 6.4).
Transcriptional control of the hrpS promoter seems to be the step directly 339 or indirectly modulated by the esaI/esaR system and this is supported by
both epistasis analysis (Table 6.2) and gene expression experiments (Fig.
6.2). In particular, post-translational activation of HrpY does not appear
to be involved in mediating the quorum sensing effects because ectopic expression of HrpY does not suppress the Hrp phenotype of the esaI strain
(Table 6.2). Addition of synthetic OHHL complemented the esaI mutation
(Fig. 6.3) but did not upregulate hrp expression of the wild-type strain at low cell densities (Fig. 6.5). Similarly, hrp expression was not upregulated at low population densities in the ∆esaR::cat strain, but it is moderately higher than the wild-type at high cell densities (Fig. 6.4).
These last observations may imply that a growth-phase-dependent regulator acts in parallel to, or downstream of, the esaR/esaI system. A similar scenario has been described for P. aeruginosa (Diggle et al.,
2002). In that species, genetic screening showed the parallel involvement of RpoS and MvaT as growth-phase regulators acting in concert with the
LasR/I and RhlR/I systems.
The observation that hrp expression in the ∆esaRI::cat strain is intermediate between the wild type and the ∆esaR::cat strain is somewhat puzzling. Assuming it is significant from a regulatory point of view, it may suggest that the OHHL produced by ∆esaI is required by other unknown AHL-receptors that also have a positive regulatory effect on hrp expression.
340 P. stewartii can multiply in plants up to ca. 105 -106 CFU g-1 tissue without causing any apparent host response (Coplin et al., 1992). After this population level is reached, a functional type III secretion system is essential for the bacteria to grow to much higher cell densities (109 CFU
g-1 and above) and cause disease symptoms (Coplin et al., 1992).
Activation of a hrp-gfpmut3[AAV] gene fusion in P. stewartii was observed
around 20-24 h post-inoculation when using low inoculum doses (M.
Merighi, unpublished observation). Other necrogenic phytobacteria of the
genera Erwinia, Pseudomonas and Xanthomonas cannot grow in plant tissue when their Hrp secretion systems are defective (Willis et al., 1991;
Bonas, 1994; Alfano and Collmer, 1997), and induction of hrp genes occurs at as little as two hours after inoculation (Boureau et al., 2003).
The fact that the Hrp pathogenicity system in P. stewartii is not only environmentally regulated (Merighi et al., 2003), but also under quorum sensing control, is consistent with the “stealth” model for growth in plant tissues. Activation of the Hrp system of P. stewartii causes the translocation of several effector proteins that are cytotoxic (Ahmad et al.,
2001; J-H, Ham and D. L. Coplin, unpublished). In other diseases, it has been shown that the host responds to this aggression with the induction of defense genes in the attempt to restrict pathogen growth (Yalpani and
Raskin, 1993; Shirasu et al., 1996; Fritig et al., 1998; Sandermann, 2000;
341 Conrath et al., 2002; Klement et al., 2003) and some pathogens even deploy active mechanisms to counteract host defenses (Abramovitch et al., 2003).
Perhaps P. stewartii delays hrp gene induction up to the point during infection when the host can no longer respond effectively to invasion. In this regard, induced defense responses have been described in corn
(Morris et al., 1998; Tiffin and Gaut, 2001), but it is not known if any of them are induced by P. stewartii. Alternatively, this delay may reflect the need for biofilm formation as a prerequisite for the interaction of the Hrp
secretion apparatus with host cells. Work in the von Bodman lab suggests
that quorum sensing controls biofilm formation on glass surfaces in P.
stewartii (Koutsoudis et al., 2002). Consequently hrp genes might be expressed only after microcolonies have already formed. This hypothesis would agree with early observations on the development of P. stewartii
populations in corn leaves (Braun, 1982). Based on Braun’s electron
microscopy studies, synthesis of EPS is completed around 12 h post-
inoculation, while the cells are still isolated or in small groups. The first
microcolonies appear only after 30 h (Braun, 1982). This timing roughly
corresponds to when hrp genes are turned on, if the plants receive low
inoculum doses. An additional advantage of expressing hrp genes in
biofilms is that the bacteria might be more resistant to hrp-induced host
oxidative responses.
342 The only other Hrp TTS system known to be regulated by quorum
sensing is P. carotovorum. This species is of particular interest because it
is evolutionarily related to P. stewartii. The RNA-binding protein RsmA
is the central regulator in this network (Fig. 6.6) and it acts by reducing
the half-life of target mRNAs. rsmA null mutants are upregulated for
exoenzyme synthesis and tissue maceration, hrpL mRNA steady-state
levels and harpinEcc production, whereas overexpression of RsmA reduces
antibiotic production and HR elicitation (Cui et al., 1995; Cui et al.,
1996; Mukherjee et al., 1996; Chatterjee et al., 2002). rsmA is controlled at the transcriptional and post-translational levels by several regulators.
The list includes the stationary phase sigma factor RpoS (Mukherjee et al., 1998) (activating rsmA), the regulatory RNA rsmB (Liu et al., 1998)
(titrating RsmA and therefore repressing its effects), the transcriptional adapter RsmC (Cui et al., 1999) (which acts negatively on rsmB and positively on rsmA expression), the two-component system GacA/GacS
(ExpA/ExpS) (activating rsmB and repressing rsmA) (Cui et al., 2001;
Hyytiainen et al., 2001), and finally by the quorum sensing system expR/expI (Andersson et al., 2000; Koiv and Mae, 2001). This means that quorum sensing controls harpin production and hrp gene expression indirectly via its effect on rsmA.
In particular, data from (Koiv and Mae, 2001) shows that an expI N- acyl-homoserine synthase knock-out mutant has increased expression of rsmA and rsmB RNAs and a net negative effect on exoenzyme production 343 and virulence. Recent work in Chatterjee’s laboratory (Chatterjee et al.,
2002) is in partial agreement with that conclusion, i.e. the absence of N-
(3-oxohexanoyl)-L-homoserine lactone (OHHL) in P. carotovorum
apparently activates transcription of rsmA but not of rsmB. The net effect of the increased concentration of cellular RsmA is the reduction of virulence and of rsmB RNA. Consequently, the effect of quorum sensing signal molecules on virulence in Pectobacterium appears to be channeled via the rsmA/B system. Work from E. Palva’s laboratory tried to define
the role for ExpR, the cognate LuxR-like receptor of OHHLs (Andersson
et al., 2000). Null expR mutations did not decrease virulence or
production of exoenzymes in vitro, but resulted in a slight increase in the
maceration capacity of the mutant strain in vivo. At the same time,
ectopic expression of ExpR reduced exoenzyme production and virulence.
These results are consistent with a model where apo-ExpR is an activator
of rsmA transcription at low OHHL concentration and increased OHHL levels may sequester ExpR, making it incompetent for DNA binding. No one has yet tested this hypothesis using biochemical approaches.
RsmA and gacA/S have been shown to be present in P. stewartii as well
(Mukherjee et al., 1998), but nothing is known about their role in quorum
sensing-mediated phenotypes and/or virulence. In spite of the similarities
in the genetic complement of the two species, substantial differences exist
between P. carotovorum and P. stewartii, both in the role of the Hrp
system in pathogenesis and the manner in which quorum sensing 344 interfaces with the Hrp regulatory cascade. The P. carotovorum Hrp
system is not required for growth in compatible hosts; tissue maceration caused by quorum-sensing controlled pectic enzymes is the primary virulence mechanism. Moreover, the effect of quorum sensing on hrp
genes is not direct. Instead, it is mediated by the global regulator RsmA,
which post-transcriptionally controls hrpL mRNA stability (Chatterjee et
al., 2002).
In P. stewartii, a necrogenic pathogen like P. syringae and Erwinia
amylovora, hrp genes are central to its virulence. In this species a direct
effect of EsaR on hrpS expression appears more likely considering that a
putative esaR box has been identified in the long 5’-UTR of hrpS
(GCCTGTCTAGTGAGGGT) (Fig. 6.7). It is interesting that the putative
esaR box is located in the IS-like element found in front of the hrpS
ribosomal binding site and that this element is absent in all the other
erwinias. Direct quorum sensing regulation of hrp genes may therefore be
a unique adaptation of P. stewartii. Other possible regulatory loops may
involve various global regulators, perhaps those described in P.
carotovorum (RsmA, rsmB, RpoS) (Fig. 6.6), but acting on hrpS. This
model is attractive because of the involvement of a known stationary-
phase regulator (RpoS) acting in parallel to the ExpI/ExpR system for
RsmA regulation (Mukherjee et al., 1998). Multiple regulators acting in
concert could be involved as well, as recently shown for P. aeruginosa
(Diggle et al., 2002). 345 6.5 SUMMARY
N-acylhomoserine lactone (HSL)-dependent quorum sensing is involved
in coordinate regulation of many bacterial phenotypes in response to
population density. In the corn pathogen P. stewartii subsp. stewartii, the
quorum sensing system encoded by the esaI OHHL synthase and esaR
receptor-regulator genes controls expression of exopolysaccharide
synthesis. In this study, we show that expression of P. stewartii hrp genes
is also under control of esaI and esaR. The expression of a hrp secretion
gene was shown to have a typical quorum sensing response curve with
time and had a threshold set at 1 x 108 CFU per ml in IM pH5.5 growth
medium. When assayed in corn whorls, an esaI mutant caused much less water-soaking than the wild-type and was unable to cause HR in tobacco.
This phenotype was suppressed by ectopic overexpression of hrpS and
hrpL, but not of hrpY. Expression of hrp secretion and effector genes was
reduced one to two orders of magnitude in an esaI mutant background and
restored in an esaRI strain. Collectively our results indicate that EsaR
mediates the effect of the esaI mutation and acts, directly or indirectly, as
a repressor of hrpS. The observations that hrp gene expression was not
upregulated significantly at low cell density in esaR mutants and that
addition of exogenous OHHL could not upregulate hrp expression in the
wild-type at low cell density suggest that additional regulation, perhaps
346 involving late exponential-stationary phase signals, may be involved. This is the first report of a Hrp type III secretion system of a stealth plant pathogen that is under direct quorum sensing control.
347
Strain or plasmid Relevant characteristics Reference
P. stewartii
SS104 Wild type, Wts+, HR+ ICPPBa
DC283 SS104 Nalr Coplin et al. (1986)
ESN51 esaI::Tn5 von Bodman et. al, 1998
ES∆R ∆esaR::cat von Bodman et al., 1998
ES∆IR ∆esaIR::cat von Bodman et al., 1998
Escherichia coli
DH10B F- mcrA ∆(mrr-hsdRMS-mcrBC) φ80dlacZ∆M15 ∆lacX74 endA1 recA1 deoR
∆(ara,leu)7697 araD139 galU galK nupG rpsL λ- Gibco BRL
Plasmids pDM2531 pPL6GUSC with a 0.9 kb BamHI fragment to create a hrpJ-uidA fusion Merighi et al., 2003
pDM2785 pPL6GUSC with a 389 bp BamHI PCR fragment to create a hrpL-uidA fusion Merighi et al., 2003
pDM2791 pPL6GUSC with a 1.1 kb HindIII-BamHI fragment from pES411 to create
a wtsE-uidA fusion Merighi et al., 2003
pMM25 pPL6GUSC with a 244 bp HindIII-BamHI PCR fragment to create a
hrpX-uidA fusion Merighi et al., 2003
pMM50 pPL6GUSC with a 1.3 kb BamHI PCR fragment to create a hrpS-uidA fusion Merighi et al., 2003
+ pMM6 pRK415 with a 0.6 kb BamHI-EcoRI hrpL PCR fragment, transcribed from Plac Merighi et al., 2003
pMM52 pRK415 with the 0.7 kb BamHI-EcoRI hrpY+ fragment from pMM46, transcribed
from Plac Merighi et al., 2003
pRF205 pVK100 with the 1.7 kb HindIII fragment containing hrpS from pES1044 Frederick et al. (1993)
+ - r r r pRK2013::Tn7 ColE1 mob traRK2 ∆ repRK2 repE kan::Tn7 (Tp , Sm , Sp ) Ditta et al. (1980)
Table 6.1. Bacterial strains and plasmids.
348
Strain Genotype Plasmid-borne hrp regulator
None hrpY hrpS hrpL
DC283 esaRI+ 2.9±0.1 a (+)b ND ND ND
ESN51 esaI- 0.8±0.4 (-) 0.6±0.25(-) 2.6±0.8(+) 2.5±0.3(+)
ES∆R ∆esaR::cat - 1.9±0.4 (+) 2.5 (+) 2.6 (+) 2.7 (+)
ES∆IR esaRI- 2.8±0.2 (+) 3.0 (+) 2.9 (+) 3.0 (+)
a Water-soaking severity in sweet corn whorls was rated using a 0 to 3 scale at 3-4
days post-inoculation. Data are the means up to 6 independent experiments with 6
replicates each ± the standard error. b HR elicitation is indicated between parenthesis.
Table 6.2. Watersoaking symptom severity ratings and hypersensitive response elicitation caused by P. stewartii esaI-esaR mutants and epistatic effects of other
Hrp regulatory genes.
349
hrpS hrpY hrpX hrpL hrpJ
GUS pMM50
GUS pMM25
pDM2531 GUS
GUS pDM2785
wtsF wtsE hrpN
GUS pDM2791 1 kb
Figure 6.1. Maps of plasmid-borne reporter gene fusions used in this study.
(Top) Map of the hrpL-hrpXY-hrpS regulatory region, including a portion of the secretion operon hrpJ . (Bottom) Map of the Hrp-secreted WtsE effector gene. The inserts cloned into the promoter probe plasmid pPL6GUSC and their orientations are shown below the maps as bars.
350
10000
1000 DC283 100 ESN51
10
GUS activity (pmol MU min-1 OD-1) 1
85 31 91 MM25 MM50 M27 M25 M27 p p D D D p p p hrp-uidA fusions
Figure 6.2. Effect of an esaI mutation on the expression of several hrp regulatory
and structural genes.
DC283 (wild-type) and ESN51 (esaI) strains carrying hrp-uidA reporter gene fusions
in plasmid pPL6GUS were grown in IM5.5 to an A600 >0.4 and GUS assays were
performed to assess hrp transcriptional activity. Data shown are the mean of two
independent experiments with two replicates each. Pair-wise averages were analyzed
using a t-test and they all were significant at P<0.05. Error bars indicate the standard
error of the mean. pMM25 is PhrpX-uidA; pMM50 is PhrpS-uidA; pDM2785 is PhrpL-
uidA; pDM2531 is PhrpJ-uidA; pDM2791 is PwtsE-uidA.
351
A600: 0.75 0.4 1.0 0.9
10000 9000 8000 7000 6000 -OHHL 5000 +10 uM OHHL 4000 3000 2000 Specific GUS activity
(pmol MU min-1 OD-1) 1000 0 DC283/pDM2531 ESN51/pDM2531
Figure 6.3. Suppression of esaI regulatory effects on hrpJ by addition of synthetic N-3-oxo-hexanoyl-homoserine lactone (OHHL).
Strains ESN51 (esaI) and DC283 (wild-type) carrying the hrpJ-uidA gene fusion in plasmid pDM2531 were grown in IM5.5 with and without the addition of 10 µM
OHHL. After 24 and 48 h of growth at 28°C, for DC283 and ESN51 respectively, the
cultures were assayed for GUS activity. Data are from one experiment with two
replicates. Optical densities (A600) of the cultures at the time of the assay are shown above the bars.
352
WT 12000 esaI esaR 10000 esaRI
8000 R2 = 0.8663 esaR 6000 R2 = 0.9403 WT 4000 R2 = 0.8341 esaIR 2000 R2 = 0.8793 GUS activity (pmol MU min-1 OD-1) esaI 0 0.01 0.1 1
OD600
Figure 6.4. Transcriptional expression of hrpJ as a function of cell density in esaI, esaR and esaIR mutants.
Cells containing a hrpJ-gusA fusion in plasmid pDM2531 were grown in LB broth
overnight, diluted 1:200 in fresh medium and grown at 28°C in flasks with shaking.
Aliquots were removed at different times for cell density and GUS activity measurements. The plot shows data from a representative experiment with 2-4 replicates per strain. Curves were interpolated by third-order polynomial regression
and the R2 of the fitting is shown. WT is strain DC283; esaI is strain ESN51 [DC283 esaI::Tn5]; esaR is strain ES∆R [DC283 ∆esaR::cat]; and esaRI is strain ES∆IR
[DC283 ∆esaIR::cat].
353
1600 DC283/pDM2531 1400 1200 1000 800 - OHHL 600 + 4 uM 400 OHHL Specific GUS activity (pmol MU min-1 OD-1) 200 0 0 0.2 0.4 0.6 OD600
Figure 6.5. Effect of addition of synthetic N-3-oxo-hexanoyl-homoserine lactone
(OHHL) on expression of hrpJ.
DC283 [wild-type] containing a hrpJ-gusA fusion in plasmid pDM2531 was grown in
LB overnight. Cells were washed in 100 mM MES pH 5.5, diluted 1:200 in IM pH5.5
medium with or without 4 µM OHHL and grown at 28°C in flasks with shaking.
Aliquots were removed at different times for measurement of cell density and GUS
activity. Data are from a representative experiment.
354
GacS/Exp expI ExpM HexA ~P RpoS
GacA/ExpA ExpR/OHHL?
rsmA rsmB RsmC
rsmB KdgR RsmA RNA
hrpL mRNA L exoenzymes σ hrp genes
Figure 6.6 Model describing the RsmA regulatory network in P. carotovorum
Refer to the Discussion section for details on the model.
355
Apo-EsaR OHHL (active)
EsaR
(inactive)
SAM +acyl- carrier protein EsaI
esa I esaR
esaR-Box ?
Stationary-phase signal?
hrpS
esaR-like box HrpY~P
HrpS BOX: GCCTGT-CTA:--GTG-AGGGT
LuxR BOX: MNCTRN-CNN:WANNGCAGNG-
EsaR BOX: GCCTGTACTA:TAGTGCA-GGT
Figure 6.7. Model for quorum sensing regulation of P. stewartii hrpS and
alignment of the putative esaR-like box (“HrpS box”).
356
CHAPTER 7
CONCLUSIONS
The proper expression of virulence factors in time and space is essential for the success of bacteria as pathogens. What molecular signals and which transduction mechanisms control plant pathogen type III secretion systems is for the most part unknown. We have seen that in P. stewartii as in most other phytobacteria, growth conditions mimicking the apoplast
(low pH, low concentration of inorganic nitrogen and phosphate sources) can activate hrp gene expression, whereas media containing complex nitrogen sources suppress it. Specific Hrp-inducing plant factors have not been characterized for any plant-microbe pathosystem. Regulation by contact has been proposed for a Hrp group II pathogen, but this is probably not a common paradigm among plant pathogens.
In the first part of this dissertation, I delineated the Hrp-activating pathway in P. stewartii (Fig. 7.1). All hrp secretion and effector genes identified in P. stewartii have “hrp boxes” in the –10/-35 region of their promoters. Genetic experiments using transcriptional fusions in both P. stewartii and E. coli demonstrated that the alternative sigma factor HrpL
357 is necessary and sufficient for recognition of “hrp box“-containing
promoters by the RNA polymerase. The expression of hrpL is controlled
by the response regulator HrpS, a receiver-less member of the NtrC family
of enhancer-binding transcription factors. A regulatory element upstream
of P. stewartii hrpS was identified by deletion analysis. These regulatory
steps are common to all group I hrp clusters in erwinias and
pseudomonads.
The most novel research presented in this dissertation is the analysis of
the two-component system HrpX/HrpY, which is also present in all
erwinias. In particular, we studied its roles in controlling hrpS and hrpL
and in autoregulation of its own operon. By using gene fusions and
ectopically expressed regulatory genes, we demonstrated that HrpY positively controls hrpS transcription and that it does not stimulate
expression of hrpL. These findings contradict a previous model published
for E. amylovora (Wei et al., 2000b), but are fully supported by the data
for P. agglomerans pv. gypsophilae, published concurrently with ours by
I. Barash’s lab (Merighi et al., 2003; Nizan-Koren et al., 2003). Sensor
and response regulators are located in the same operon, which is also
autoregulated by HrpY. We further showed that a conserved Asp residue
is required for the activity of HrpY. The response regulator was purified
as a recombinant protein for in vitro studies. We demonstrated that HrpY
is phosphorylated in vitro at the D57 residue and we confirmed its direct
interaction with its target promoters by gel shift assays and preliminary 358 DNA footprinting analysis. In these studies, phosphorylation appeared to increase the affinity of HrpY for the hrpS promoter (Fig. 7.1). However,
phosphorylation may not be needed for activation of the RNA
polymerase/DNA complex, but simply for increased oligomerization and
cooperative binding at multiple regulatory sites within the hrpS promoter
region.
Regulation of the hrpS promoter probably involves more factors than
just HrpY. This was suggested by the following observations: (a) Crude
extracts from a P. stewartii hrpY null mutant still retard hrpS promoter
fragments in gel shift experiments. (b) An IHF-like site has been
identified several hundred bp upstream of the promoter of hrpS. (c)
Deletion analysis of hrpS indicates the existence of an operator for an
unknown repressor (and in this respect, HrpY may function as an
antirepressor). (d) The quorum-sensing regulator EsaR was shown to
mediate the repression of hrpS in an OHHL-synthase mutant and a
potential EsaR-like box was found downstream of the hrpS promoter. (f)
hrpS expression is upregulated by a mutation in a nikR-like gene (Ham
and Coplin, unpublished).
The critical role of regulation at the hrpS promoter in integrating
multiple regulatory pathways is also highlighted by the results described
in Chapter 5. The environmental and metabolic signals generated upon
growth in media with high pH, high osmolarity, nicotinic acid, complex
nitrogen sources and some organic acids as sole C sources all repress 359 hrpS. Moreover, only a few of these (nitrogen sources, citrate and
succinate) are mediated by HrpX /HrpY. Therefore, HrpX may function
primarily as a metabolic starvation sensor. Its predicted cytoplasmic
location, the presence of PAS domains in its input region and the essential
role of these domains in hrp gene activation all support this hypothesis.
The remaining external signals must be transduced via alternative
regulatory pathways acting on PhrpS. For example, in enteric bacteria, osmolarity and pH are usually sensed by two-component systems or by direct changes in the topology of the promoter DNA (Finlay and Falkow,
1997). Given that HrpY is absolutely required for virulence and expression of hrpS, most of these postulated parallel regulatory mechanisms probably involve repression (Fig. 7.1).
The presence of so many independent mechanisms controlling a single master regulator may reflect the need to tightly regulate hrp gene transcription, allowing full expression only upon the fulfillment of multiple prerequisites in addition to simple starvation. Our discovery that the expression of hrpS is indirectly autoregulated by a novel transcriptional strategy further stresses the importance of maintaining a stable level of HrpS once the Hrp-pathway has been activated by external clues. The need for positive autoregulation perhaps also reflects high turnover rates for HrpS. This possibility is supported by the observation
360 that the Lon protease is a negative regulator of hrp gene expression in P.
stewartii, similar to what was described for P. syringae (Bretz et al.,
2002).
In addition to the primary role of HrpY in activating hrpS, we found
that, when it is overexpressed, HrpY has a negative effect on the basal
expression of hrpL from its tandem σ 70 promoter. This has not been characterized or described in any other erwinias. We still need to show that this repression happens in P. stewartii but it could represent a novel means to directly downregulate a critical regulator of hrp/hrc genes in order to dampen the autoregulatory loop triggered by HrpS via the hrpL promoter.
Perhaps my most surprising finding was the discovery that hrp genes are also regulated by quorum sensing, again at the level of hrpS transcription
(Fig. 7.1). Regulation by quorum sensing implies that hrp genes are only fully expressed when a local quorum of cells is reached (i.e. after microcolonies are formed). However, it is believed that the early multiplication in planta of most necrogenic phytobacteria is dependent on early induction of hrp genes. This tautology may be explained by the ability of P. stewartii to grow in plant tissues to considerable population densities as endophyte independently of hrp genes. In other words, hrp genes are used in this species only for a short, but critical, final attack.
The reasons for this strategy are unknown, but it could be that a quorum
361 sensing-dependent protective biofilm must be formed before the pathogen starts secreting effectors that could trigger host defenses.
Alternatively, or concurrently, P. stewartii may use a “brute-force” approach, delaying the critical attack of the host to when a sufficient population of invading cells is reached, and therefore overwhelming host defenses.
362
High pH, High osmolarity Nicotinic acid Nitrogen starvation? EsaR
HrpY~P σL ~P HrpX Hrp regulon
High KD Low KD High KD
UAS
(+/-)?
IHF σ54
Lon HrpS
Figure 7.1. Global regulation of hrp genes in P. stewartii.
363
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