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REGULATION OF THE RNase E-MEDIATED TURNOVER OF NON-CODING sRNAs CsrB AND CsrC
By
YUANYUAN LENG
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
2017
© 2017 Yuanyuan Leng
To my family and husband
4
ACKNOWLEDGMENTS
I want to express my deepest gratitude to my advisor, Dr. Tony Romeo for giving
me the opportunity to work in his lab and for his guidance, encouragement, care and
support during my PhD study. Under his excellent guidance, I have not only built good
skills and expertise in different areas, but also learned how to be a good scientist, to
think critically and independently, and to keep myself enthusiastic about research. I
truely appreciate his help to develop my presentation skills and his support during my
job search. I would also like to thank my committee members, Dr. Julie Maupin-Furlow,
Dr. Maurice Swanson, Dr. Wayne L. Nicholson and Dr. James Preston for their valuable
suggestions and comments in my research.
I am grateful to my colleagues in Dr. Romeo’s lab, especially Dr. Christopher
Vakulskas for his help and guidance when I joined the lab. I am also thankful to
Anastasia Potts for collaborating with me on the work in chapter 4 and for her help with
the RNA-seq library preparation and RNA-seq data analysis, to Dr. Archana Pannuri for
performing Northern blots to determine the effect of enolase on CsrB/C turnover in
chapter 3, to Dr. Tesfelam Zere for his help with Phos-tag gel, and to John Rice for his
help with lab supplies ordering. They have given me much help in my research studies
and a lot of smiles in my lab life. I want to thank Dr. Sixue Chen, Dr. Jin Koh and
Fanchao Zhu at the ICBR Proteomics and Mass Spectrometry Core for their support
with my research. I would like to thank my parents, who provide me endless support to
study abroad and encourage me when I encounter frustrations in work and life. Most
importantly, I wish to thank my loving and supportive husband, Likui Feng, for providing
unending inspiration and for always being there to help me.
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TABLE OF CONTENTS
page
ACKNOWLEDGMENTS .................................................................................................. 4
LIST OF TABLES ............................................................................................................ 8
LIST OF FIGURES .......................................................................................................... 9
ABSTRACT ................................................................................................................... 11
CHAPTER
1 GENERAL INTRODUCTION .................................................................................. 13
Bacterial Gene Expression Regulation ................................................................... 13 Bacterial Transcriptional Regulation ................................................................. 13
Bacterial Posttranscriptional Regulation ........................................................... 14 Bacterial small regulatory RNAs ................................................................ 15 RNA binding proteins involved in gene expression regulation ................... 16
Turnover of sRNAs ................................................................................................. 17 Turnover of Base-pairing sRNAs ...................................................................... 18
Turnover of Protein Binding sRNAs .................................................................. 20 The Carbon Storage Regulator (Csr) System ......................................................... 21
CsrA/RsmA ....................................................................................................... 21
Regulation of the Csr System ........................................................................... 22
Transcription of CsrA Inhibitory sRNAs ............................................................ 23 Regulation of the Turnover of CsrA Inhibitory sRNAs ...................................... 24 CsrA Regulates its Own Expression ................................................................. 25
Interaction of Csr System with Other Regulatory Pathways ............................. 25 Project Rationale and Objectives ............................................................................ 27
2 REGULATION OF CsrB/C sRNA DECAY BY EIIAGlc ............................................. 30
Introduction ............................................................................................................. 30 Materials and Methods............................................................................................ 33
Bacterial Strains and Culture Conditions .......................................................... 33 Construction of Plasmids and Mutant Strains ................................................... 33 Expression and Purification of EIIAGlc and CsrD Variants ................................ 35
Northern Blotting .............................................................................................. 35 Western Blotting ............................................................................................... 36
Protein Pull-down Assays ................................................................................. 37 Determination of the EIIAGlc Phosphorylation State .......................................... 38 Gel Filtration Analysis of EIIAGlc in Complex with the EAL Domain or Other
CsrD Variants ................................................................................................ 38 Results .................................................................................................................... 39
EIIAGlc Activates CsrB/C Decay via CsrD ......................................................... 39
6
In vitro Binding of EIIAGlc to CsrD Requires the EAL Domain ........................... 40
EIIAGlc Regulates CsrB/C Decay in a Phosphorylation-dependent Manner ...... 42
cAMP-Crp Modestly Represses CsrB Turnover ............................................... 43 CsrB/C Decay is Regulated in Response to Carbon Availability via the
Phosphorylation State of EIIAGlc .................................................................... 44 EIIAGlc and MshH Promote Csr sRNA Decay in Vibrio cholerae ....................... 45
Discussion .............................................................................................................. 46
3 EXPLORING THE MOLECULAR MECHANISM BY WHICH CsrD FACILITATES CsrB/C TURNOVER........................................................................ 61
Introduction ............................................................................................................. 61 Materials and Methods............................................................................................ 63
Media and Growth Conditions .......................................................................... 63
Construction of Strains and Plasmids ............................................................... 64 Gel Mobility Shift Assay .................................................................................... 64 In vitro RNase E Cleavage Assays ................................................................... 65
Affinity Purification of CsrD and its Binding RNAs ............................................ 65 Affinity Purification of in vivo Synthesized CsrB and its Associated Proteins ... 66
Mass Spectrometry .......................................................................................... 67 Protein Purification and Western Blots ............................................................. 69
RNA Purification and Northern Blots ................................................................ 69 Results and Discussion........................................................................................... 69
EIIAGlc is not Capable to Stimulate the Binding of CsrD to CsrB in vitro ........... 69 CsrA Influences the CsrD-CsrB Interaction in vitro ........................................... 70
CsrD cannot Facilitate CsrA-mediated Protection of CsrB Cleaved by RNase E ........................................................................................................ 70
CsrD is Unlikely to Bind CsrB in vivo ................................................................ 71
CsrB Associated Proteins and their Influence on CsrB Turnover ..................... 72 Conclusion .............................................................................................................. 75
4 EPISTASIS ANALYSIS USING RNA-SEQ (EPI-SEQ) TO EXPLORE THE REGULATORY ROLE OF CsrD ............................................................................. 84
Introduction ............................................................................................................. 84
Materials and Methods............................................................................................ 86 Media and Growth Conditions .......................................................................... 86 Construction of Strains and Plasmids ............................................................... 87 Glycogen Biosynthesis ..................................................................................... 87
RNA Extraction and Purification ....................................................................... 87 Northern Blotting .............................................................................................. 88 RNA-seq Library Preparation ........................................................................... 88
RNA-seq Data Analysis .................................................................................... 88 qRT-PCR .......................................................................................................... 89
Results and Discussion........................................................................................... 89 Construction and Characterization of Bacterial Strains .................................... 89
7
CsrA Retains its Global Role in Regulating Transcript Levels in the Absence of CsrD .......................................................................................................... 91
CsrD Effects on Gene Expression Require CsrA ............................................. 93 CsrB/C Have Strong Effects on Gene Expression in the Absence of CsrD ...... 94 CsrD Regulates the Majority of its Target Genes in a CsrB/C Dependent
Manner .......................................................................................................... 94 Conclusion .............................................................................................................. 96
5 GENERAL DISCUSSION AND FUTURE PERSPECTIVES ................................. 104
APPENDIX
A SUPPLEMENTARY FIGURES ............................................................................. 109
B SUPPLEMENTARY TABLES ............................................................................... 110
LIST OF REFERENCES ............................................................................................. 124
BIOGRAPHICAL SKETCH .......................................................................................... 141
8
LIST OF TABLES
Table page 2-1 Molecular weight of CsrD in solution .................................................................. 54
3-1 Proteins co-purifying with Strepto-CsrB identified from band A by mass-spectrometry ....................................................................................................... 82
3-2 Proteins co-purifying with Strepto-CsrB identified from band B by mass-spectrometry ....................................................................................................... 83
B-1 Bacterial Strains ............................................................................................... 110
B-2 Plasmids used in this study. ............................................................................. 113
B-3 Primers used in this study. ............................................................................... 115
B-4 Transcription regulators regulated by CsrA ...................................................... 118
B-5 sRNAs regulated by CsrA at the transcript level ............................................... 123
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LIST OF FIGURES
Figure page 1-1 Outline of the Csr system in E. coli (Vakulskas et al., 2015) .............................. 29
2-1 EIIAGlc affects CsrB/C decay rates and levels in E. coli ...................................... 50
2-2 EIIAGlc (crr) activates CsrB/C decay via CsrD, but does not enhance cellular CsrD levels ......................................................................................................... 51
2-3 EIIAGlc interacts specifically with the EAL domain of CsrD. ................................ 52
2-4 Gel filtration chromatography of CsrD variants ................................................... 53
2-5 CsrD binds only to unphosphorylated EIIAGlc in pull-down assays. .................... 54
2-6 Effects of cAMP-Crp on CsrB/C decay. .............................................................. 55
2-7 Effects of carbon sources on EIIAGlc phosphorylation and CsrB/C decay in minimal media .................................................................................................... 56
2-8 The phosphorylation state of EIIAGlc before and 10 min after shift from LB broth into minimal media. ................................................................................... 57
2-9 CsrB/C decay rates and levels after shift from LB to minimal media .................. 58
2-10 EIIAGlc and MshH (CsrD ortholog) affect CsrB and CsrD decay in V. cholerae.. ............................................................................................................ 59
2-11 Proposed model for the effect of carbon availability on CsrB/C decay. .............. 60
3-1 CsrB decay is regulated in response to carbon availability through its effect on CsrA and CsrD antagonism (Vakulskas et al., 2016).. ................................... 76
3-2 EIIAGlc has no effect on the binding of CsrD to CsrB RNA in vitro.. .................... 77
3-3 CsrA influences the CsrD-CsrB complex in vitro.. .............................................. 78
3-4 Effect of EIIAGlc and CsrD on RNase E-dependent cleavage of CsrB in vitro.. ... 79
3-5 CsrB is not copurified with FLAG-tagged CsrD. .................................................. 80
3-6 CsrD is not recovered by Strepto-CsrB in vivo ................................................... 81
3-7 Enolase has little or no effect on CsrB turnover.................................................. 83
4-1 Two proposed model depicting the epistatic relationship between CsrA and CsrD in regulating mRNA abundance.. ............................................................... 97
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4-2 Properties of bacterial strains used in this study.. ............................................... 98
4-3 CsrA retains its global role in regulating mRNA levels in the absence of CsrD. 100
4-4 Venn diagram depicting the overlap of differentially expressed genes induced by csrD deletion/overexpression in csrA WT and csrA mutant backgrounds.. .. 102
4-5 Venn diagram depicting the overlap of differentially expressed genes induced by csrB/C deletion or csrB overexpression in csrD WT and csrD mutant backgrounds. .................................................................................................... 102
4-6 CsrD regulate the majority of its target genes in a CsrB/C dependent manner. 103
A-1 EIIAGlc stimulates CsrB decay in the MG1655 csrDFLAG strain. ......................... 109
A-2 Decay of CsrB/C in a strain expressing EIIAGlc H91D is similar to that of EIIAGlc H91A ..................................................................................................... 109
A-3 CRP has minimal or no effect on CsrD protein levels. ...................................... 109
11
Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy
REGULATION OF THE RNase E-MEDIATED TURNOVER OF NON-CODING sRNAs
CsrB AND CsrC
By
Yuanyuan Leng
August 2017
Chair: Tony Romeo Major: Microbiology and Cell Science
Bacteria utilize complex regulatory system to coordinate their gene expression
and make life style decisions in response to changing environment. Carbon Storage
Regulator (Csr) is a conserved global regulatory system in Gammaproteobacteria and it
uses the sequence-specific RNA-binding protein CsrA to activate or repress gene
expression at the posttranscriptional level. In Escherichia coli, CsrA activity is regulated
by two non-coding sRNAs, CsrB and CsrC, which bind to multiple CsrA dimers and
thereby sequester this protein away from its mRNA targets. Both the synthesis and
turnover of CsrB/C are regulated and play important roles in determining the steady
state levels of CsrB/C. Their turnover is initiated with RNase E cleavage, and the decay
intermediates are subsequently degraded by PNPase. This RNase E-mediated turnover
of CsrB/C requires CsrD protein although the exact molecular mechanism remains
unclear. In this study, we revealed the physiological role of CsrD in coupling CsrB/C
sRNA decay to the availability of a preferred carbon source. We demonstrated that
EIIAGlc of the glucose-specific PTS system regulates CsrB/C turnover in a
phosphorylation dependent manner and only the unphosphorylated form of EIIAGlc binds
to CsrD and is capable of activating CsrB/C turnover. On the other hand, the
12
phosphorylated form of EIIAGlc indirectly and modestly represses CsrB turnover via
cAMP-Crp. In addition, we provided evidence that although CsrD binds to CsrB in vitro,
it does not appear to bind CsrB in vivo and it fails to facilitate the cleavage of CsrA-
protected CsrB by RNase E in vitro. These data suggest that CsrD might act indirectly
and require other unknown factor(s) to facilitate the CsrB/C turnover. Furthermore, the
global regulatory role of CsrD on gene expression was explored on a genome-wide
scale using epistasis analysis and RNA-seq. The transcriptomic data indicated that
CsrD mediates changes in gene expression primarily through its effects on CsrB/C
stability and thereby CsrA activity. Our data also implied that CsrD affects expression of
some genes through an alternative pathway independent of CsrB/C, suggestive of other
regulatory role(s) of CsrD in addition to affecting CsrB/C turnover.
13
CHAPTER 1 GENERAL INTRODUCTION
Bacterial Gene Expression Regulation
Bacteria are constantly exposed to a variety of environmental stresses, such as
changes in temperature, pH, osmolarity and nutrient availability. To survive in the
changing environments, bacteria have developed complex regulatory systems to
coordinate their gene expression in response to cellular and environmental stimuli and
properly adjust cellular physiology and metabolism.
Bacterial Transcriptional Regulation
Expression of genes can be regulated at the transcriptional level by modulating
transcription initiation with alternative sigma factors (σ factors), transcription factors, or
small ligands (Browning & Busby, 2004, Balleza et al., 2009). Under stress conditions,
alternative σ factors, subunits of RNA polymerase (RNAP) holoenzyme, direct RNAP to
initiate transcription of specific gene sets related to stress responses. The best-studied
alternative σ factor in Escherichia coli is RpoS (Weber et al., 2005). It accumulates in
the cells entering stationary phase or during starvation and controls expression of up to
10% of the E. coli genes (Weber et al., 2005, Patten et al., 2004). This thereby triggers
a global stress response and allows the bacterial cells resistant to stress conditions.
Modulation of transcriptional gene expression during environmental shifts is also
mediated by a large set of transcription factors in bacteria (Browning & Busby, 2004).
By directly binding to promoters of target genes, transcription factors either activate or
repress target gene expression. E. coli possesses seven global transcription factors,
CRP, Fis, ArcA, Lrp, FNR, IHF and NarL, which control expression of numerous genes
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involved in complex cellular processes, whereas most other transcription factors govern
only a single promoter.
Small ligands provide another mechanism by which cells ensure proper gene
expression following external stimuli. (p)ppGpp, cyclic AMP (cAMP) and cyclic di-GMP
(c-di-GMP) are three common and versatile small ligands that bacteria utilize to control
various important biological processes, including motility, biofilm formation, carbon
transport and metabolism, virulence and stringent response (Battesti et al., 2011, Jenal
et al., 2017, Römling et al., 2013, Botsford & Harman, 1992). The concentrations of
these ligands fluctuate in response to different environmental stimuli, and they function
by modulating activity of their binding proteins, such as RNAP (ppGpp) (Dalebroux &
Swanson, 2012), CRP (cAMP) (Fic et al., 2009), effector proteins or riboswitches (c-di-
GMP) (Sudarsan et al., 2008, Hengge, 2009).
Bacterial Posttranscriptional Regulation
In addition to be regulated at the transcriptional level, bacterial gene expression
is controlled at the posttranscriptional level. In recent years, a number of new small
regulatory RNAs (sRNAs) and RNA-binding proteins (RBPs) have been identified in
bacteria as major posttranscriptional regulators (Gottesman & Storz, 2011, Storz et al.,
2011, Van Assche et al., 2015). These posttranscriptional regulators typically regulate
gene expression by altering translation initiation, transcription elongation and/or
transcript stability. In a few cases, mRNAs have also been described to have regulatory
functions (Dugar et al., 2016, Sterzenbach et al., 2013). For example, the leader region
of flaA mRNA encoding the major fimbriae in Campylobacter jejuni, acts as an mRNA-
derived RNA antagonist of CsrA, which is the global posttranscriptional regulatory
protein (Dugar et al., 2016).
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Bacterial small regulatory RNAs
Bacterial sRNAs range in length from approximately 50 to 500 nucleotides and
they play critical regulatory roles in almost every aspect of bacterial physiology, from
metabolism, virulence, biofilm formation to outer membrane synthesis (Gottesman &
Storz, 2011, Storz et al., 2011). The expression of bacterial sRNAs is often induced
under specific stress conditions, such as glucose starvation, glucose-phosphate stress,
iron deficiency and oxidative stress, and the mechanism of action of these sRNAs has
been extensively studied in recent years.
The majority of sRNAs act by base pairing with their target mRNAs with either
extended or limited complementarity. Pairing between sRNA and mRNA is often
incomplete and requires the presence of the RNA chaperone protein Hfq, which
facilitates base pairing by stabilizing sRNA-mRNA duplexes or affecting the secondary
structure of sRNA or mRNA (Vogel & Luisi, 2011). sRNAs mostly act as a negative
regulator of gene expression by blocking ribosome binding, which in turn inhibits
translation initiation, or by recruiting RNases to destabilize target mRNAs. In some
cases, sRNAs activate expression of target mRNAs by blocking mRNA cleavage or by
disrupting an inhibitory secondary structure that occludes ribosome binding.
In other cases, sRNAs exert their functions by mimicking the nucleic acid
substrates of proteins and thereby sequestering protein activity. So far, only a few
sRNAs of this class have been identified and characterized in bacteria. E. coli 6S RNA
contains a large central “loop” flanked by long double helical arms which mimics a open
DNA promoter complex. Thus, 6S RNA can bind tightly to the housekeeping form of
RNAP (σ70-RNAP) and inhibits its activity, leading to down-regulation of a subset of σ70–
dependent promoters (Wurm et al., 2010, Burenina et al., 2015). Another well known
16
example is Csr/Rsm sRNAs, which contain many GGA sequences, allowing them to
bind CsrA/RsmA with high affinity, sequestering CsrA/RsmA from lower affinity target
mRNAs and antagonizing their activities (Fig. 1-1) (Romeo et al., 2013).
RNA binding proteins involved in gene expression regulation
Bacterial RNA binding proteins bind to target mRNAs and posttranscriptionally
regulate their expression by altering translation initiation, stability, and/or transcript
elongation. They exert their regulatory effects by modulating 1) the susceptibility of
mRNAs and/or sRNAs to RNases, 2) ribosome binding, 3) interaction of mRNA targets
with sRNA or proteins, and 4) formation of transcription terminator/antiterminator (Van
Assche et al., 2015, Vogel & Luisi, 2011, Romeo et al., 2013).
Two representative posttranscriptional regulatory proteins in bacteria are Hfq and
CsrA. Hfq is a widespread bacterial protein that resembles the eukaryotic Sm family of
proteins involved in splicing and mRNA degradation (Brennan & Link, 2007). Hfq forms
a stable ring-like, multimeric quaternary architecture that supports its interaction with
target RNAs. It affects translation initiation and transcript stability of target mRNAs by
facilitating the formation of stable sRNA-mRNA duplexes, such as Spot42-galK (Møller
et al., 2002), MicA-ompA (Johansen et al., 2006, Udekwu et al., 2005), and SgrS-ptsG
(Kawamoto et al., 2006). Thus, Hfq plays important roles in a variety of physiological
processes, from catabolite repression, envelope stress, biofilm formation, motility, metal
homeostasis to virulence (Holmqvist et al., 2016). CsrA is another major
posttranscriptional regulatory protein that contributes to complex posttranscriptional
networks (Romeo et al., 2013, Vakulskas et al., 2015). Its mechanism of action,
regulation and physiological roles will be discussed in a later section.
17
In addition to Hfq and CsrA, other RNA binding proteins have been identified to
regulate expression of target mRNAs using distinct mechanisms, including RNA
helicases (Kaberdin & Bläsi, 2013, Vakulskas et al., 2014), ribonucleases (Saramago et
al., 2014), cold shock proteins CspA and CspE (Barria et al., 2013, Michaux et al.,
2017), S1 protein (Hajnsdorf & Boni, 2012), adaptor protein RapZ (Göpel et al., 2013),
and RNA chaperone ProQ (Smirnov et al., 2016, Chaulk et al., 2011). With the constant
development of biochemical techniques for the probing of direct protein-DNA/RNA
interactions, there is no doubt that new posttranscriptional regulatory RBPs will be
identified in the near future.
Turnover of sRNAs
sRNA levels must be tightly controlled to govern expression of target mRNAs or
activity of target proteins. Expression of many sRNA is extensively regulated by
transcription regulators, including global transcription factors Lrp, Crp and Fur (Chaulk
et al., 2011, Papenfort & Vogel, 2011, Thomason et al., 2012, Massé & Gottesman,
2002), σ factors RpoE and RpoS (Opdyke et al., 2004, Guo et al., 2014), two
component signal transduction systems (TCS) (Suzuki et al., 2002, Mandin &
Gottesman, 2010), and stringent response components (p)ppGpp and DksA (Edwards
et al., 2011). This extensive regulation ensures bacterial cells fine-tune sRNA levels and
global gene expression in response to different environmental stimuli. In recent years,
the turnover pathway of sRNAs has also been revealed to play important roles in
determing the steady state levels of sRNA (Saramago et al., 2014, Andrade et al.,
2012). Here, the RNases and RBPs involved in sRNA turnover are reviewed.
18
Turnover of Base-pairing sRNAs
Pairing of sRNA and target mRNA often induces coupled degradation of the
sRNA-mRNA duplex, which mainly involves cleavage by RNase E and RNase III
(Saramago et al., 2014). RNase E is an essential endoribonuclease in Gram-negative
bacteria, which is responsible for bulk RNA turnover, and it prefers to cleave single-
stranded AU rich regions with adjacent stem-loop structures (Górna et al., 2012,
Mackie, 2013). It provides the scaffolding core of the RNA degradosome protein
complex, which also contains polynucleotide phosphorylase (PNPase), RNA helicase B
(RhlB), and the glycolytic enzyme enolase. RNase E generally recognizes substrates by
two mechansims (Mackie, 2013). The first one requires a 5’ monophosphate group on
the target transcript. By binding to the RNA 5’ end, the affinity of RNase E for the
transcript increases, which allostericall activates the activity of RNase E and facilitates
further cleavage of the transcript to the 3’ end (Prévost et al., 2011, Bandyra et al.,
2012). The other mechanism that RNase E utilizes is termed ‘direct entry’ and it
bypasses the 5’ end requirement. In this mechanism, RNase E binds close to the
internal cleavage site and cleaves the transcript (Baker & Mackie, 2003). RNAs that are
free of protein binding are more susceptible to the ‘direct entry’ cleavage (Göpel et al.,
2013). RNase E often acts together with Hfq to trigger the coupled degradation of
sRNA-mRNA duplexes (Basineni et al., 2009). Mostly, Hfq triggers RNA degradation by
recruiting RNase E and PNPase for cleavage (Sharma et al., 2007, Massé et al., 2003,
Göpel et al., 2013, Vogel & Luisi, 2011, Mohanty et al., 2004, Hankins et al., 2010). In
some cases, Hfq blocks the degradation of sRNA-mRNA duplex by competing with
RNase E for RNA binding (Moll et al., 2003). RNase E also degrades specific duplexes
independent of Hfq (Andrade et al., 2012, Lee & Groisman, 2010, Göpel et al., 2013). In
19
the absence of Hfq, RNase E can be recruited by other adapter proteins to the target
sRNA (Göpel et al., 2013). For example, E. coli sRNA GlmZ binds and activates glmS
mRNA, which encodes glucosamine-6-phosphate (GlcN6P) synthase. The RNase E-
mediated degradation of GlmZ requires a specialized adaptor protein, RapZ. Under low
levels of GlcN6P, sRNA GlmY accumulates and antagonizes RapZ activity. This results
in stabilization of GlmZ and thereby synthesis of GlcN6P (Göpel et al., 2013).
RNase III is another important endoribonuclease responsible for the degradation
of paired sRNA-mRNA (Afonyushkin et al., 2005, Deltcheva et al., 2011). It specifically
recognizes and cleaves double-stranded (ds) RNA, and is able to cleave many sRNAs
in complex with their target mRNAs. For example, in Salmonella enterica serovar
Typhimurium, RNase III cleaves MicA, the negative regulator of outer membrane porins,
when bound to its target ompA mRNA (Viegas et al., 2011).
In a few cases, binding of sRNA to mRNA only promotes cleavage of mRNA but
not sRNA. One example is sRNA MicC. After degradation of its target ompD mRNA by
RNase E, MicC is released from the duplex and is still capable of pairing and inducing
cleavage of another ompD transcript (Bandyra et al., 2012).
Many sRNAs that are free of binding to the RNA chaperone Hfq and their target
mRNAs exist transiently in the cell. In addition to RNase E (Andrade et al., 2012, Lee &
Groisman, 2010, Göpel et al., 2013), PNPase is another key factor responsible for their
degradation (Andrade et al., 2012). When not associated with Hfq, the U rich region of
the 3’ end of sRNA is exposed and can be recognized by PNPase for rapid degradation
(Sauer & Weichenrieder, 2011, Andrade et al., 2012). PNPase was observed to make a
greater contribution than RNase E in degrading many of the Hfq-free sRNAs, including
20
MicA, GlmY, RyhB, and SgrS, especially in stationary phase cells (Andrade et al.,
2012).
RNase J1/J2 and RNase Y are major factors in the processing and turnover of
RNAs in Gram-positive bacteria. Similar to RNase E, RNase J1/J2 specifically
recognize an AU-rich single-stranded RNA segments and prefer monophosphorylated 5’
ends (Even et al., 2005, Deikus & Bechhofer, 2009). However, their target sRNas have
not been explored much. A few sRNAs, such as RatA in Bacillus subtilis (Commichau &
Stülke, 2012) and RsaA and Sau63 in Staphylococcus aureus (Abu-Qatouseh et al.,
2010, Geissmann et al., 2009), are identified to be cleaved by RNase Y.
Turnover of Protein Binding sRNAs
Studies on the degradation of CsrA/RsmA inhibitory sRNAs have been limited to
only a few species. E. coli CsrB/C turnover is mediated by RNase E and PNPase and
facilitated by CsrD protein (Suzuki et al., 2006). This process will be discussed in a later
section.
6S RNA functions as a global transcription regulator by interacting with σ70 –
RNAP and inhibiting its activity (Wassarman & Storz, 2000). A recent study has
revealed that E. coli 6S RNA is cleaved by an RNase Z family protein, RNase BN, which
acts in maturation of tRNA precursors (Dutta & Deutscher, 2010, Dutta et al., 2012).
RNase BN level is growth phase dependent, it reaches the highest level in early
exponential phase, and is essentially absent in stationary phase. This leads to low level
of 6S in exponential phase and accumulation of 6S as entering stationary phase (Chen
et al., 2016). As a consequence, the housekeeping RNA polymerase is available for
enhanced transcription during exponential phase of growth, while under stationary
21
phase, its activity is repressed by accumulated 6S RNA, resulting in decreased
expression of σ70-dependent transcription.
The Carbon Storage Regulator (Csr) System
CsrA/RsmA
To cope with changes in nutrient availability, bacteria utilize both transcriptional
and posttranscriptional regulators to modulate gene expression. RpoS, cAMP-Crp and
ppGpp were discussed earlier as transcriptional regulators governing gene expression
under nutrient-limited conditions. In addition, CsrA and its orthologs RsmA and RsmE
posttranscriptionally modulate gene expression and serve as major switches of bacterial
lifestyle between rapid growth and stress resistant growth. CsrA/RsmA is conserved
throughout Gammaproteobacteria and globally represses stationary phase processes,
including gluconeogenesis, glycogen synthesis, stringent response, biofilm formation
and quorum sensing, while it stimulates expression of genes related to motility and
glycolysis for rapid growth (Romeo, 1998, Babitzke & Romeo, 2007, Romeo et al.,
2013, Vakulskas et al., 2015).
CsrA/RsmA regulates gene expression via a direct binding to mRNA targets.
Structural studies of E. coli CsrA, its homolog RsmA in Yersinia enterocolitica, and the
CsrA/RsmA in complex with RNA revealed that CsrA/RsmA is homodimeric protein with
two RNA binding surfaces formed primarily by the parallel β-1 and β-5 strands of
oppossing polypeptides (Gutiérrez et al., 2005, Heeb et al., 2006, Mercante et al., 2006,
Schubert et al., 2007). The two RNA binding surfaces of CsrA/RsmA allows it to bridege
tow target sites on a single RNA molecule (Mercante et al., 2009). In vitro selection and
in vivo UV crosslinking with RNA deep sequencing (CLIP-seq) revealed that CsrA
22
prefers to recognize AUGGA sequences present in apical loops of hairpin structures in
many bacterial species (Dubey et al., 2005, Holmqvist et al., 2016).
To achieve its regulatory role, CsrA often interacts with the 5’-UTR or early
coding region and modulates translation efficiency (Baker et al., 2002, Baker et al.,
2007, Dubey et al., 2003), transcript stability (Liu et al., 1995, Wang et al., 2005,
Yakhnin et al., 2013) and/or transcription elongation (Figueroa-Bossi et al., 2014) (Fig.
1-1). CsrA binding mostly blocks expression, but it also works as a positive regulator of
gene expression.
Regulation of the Csr System
In Gammaproteobacteria, CsrA/RsmA activity is primarily controlled through
sequestration by non-coding sRNAs, such as CsrB/C in E. coli. These sRNAs contain
multiple stem-loops with the conserved CsrA binding GGA motifs, which allow them to
bind CsrA with high affinity, sequester CsrA away for lower affinity mRNA targets and
therefore antagonize CsrA activity (Fig. 1-1) (Liu et al., 1997). In E. coli, CsrB appears
to be the principal sRNA antagonizing CsrA activity under laboratory growth conditions
(Weilbacher et al., 2003, Liu et al., 1997). Deletion of CsrB affects expression of a
number of CsrA target genes and several physiology processes (Liu et al., 1997). The
effects of CsrC on gene expression and celluar physiology are not observed unless
csrB is deleted (Weilbacher et al., 2003). McaS sRNA is recently identified as another
CsrA inhibitory regulator, but its effects are only substantial when overexpressed
(Jørgensen et al., 2013). The redundant effect of these antagonizing sRNAs might
enhance the robustness of the Csr regulation, and also ensure fine-tuning of CsrA
activity in response to different stress conditions.
23
Despite the widespread distribution of CsrA, a few bacteria, like Gram-negative
Legionella pneumophila and Gram-positive Bacillus subtilis, appear to lack the CsrA
inhibitory sRNAs. In these species, CsrA activity is regulated by a protein antagonist,
FliW, and these proteins together with a cytoplasmic Hag (flagellin) protein contribute to
the tight control of flagellin homeostasis (Mukherjee et al., 2011, Dugar et al., 2016).
Additionally, two mRNAs (flaA and fimAICDHF) derived RNA antagonists of CsrA inhibit
CsrA activity when highly expressed (Dugar et al., 2016, Sterzenbach et al., 2013).
In bacteria that possess the CsrA inhibitory sRNAs, fluctuations in the levels of
these sRNAs play a central role in regulating the Csr system (Weilbacher et al., 2003,
Liu et al., 1997). Their synthesis and stability have been studied in a few bacteria
species (Vakulskas et al., 2015) .
Transcription of CsrA Inhibitory sRNAs
Multiple factors ensure appropriate expression of Csr sRNAs under different
environmental signals. Transcription of E. coli CsrB/C is primarily regulated by the BarA-
UvrY two component signaling transduction system (TCS), which is highly conserved
across Gammaproteobacteria (Suzuki et al., 2002, Chavez et al., 2010, Martínez et al.,
2014, Zere et al., 2015). BarA protein is a membrane-bound sensor-kinase and its
activity is activated by acetate, formate and other carboxylate compounds (Chavez et
al., 2010, Huang et al., 2008). The response regulator UvrY is subsequently
phosphorylated and stimulates CsrB/C transcription (Zere et al., 2015). In a complex
negative feedback loop, CsrA regulates csrB/C transcription by activating both the
expression of the response regulator UvrY and its phosphorylation by the sensor-kinase
BarA (Suzuki et al., 2002, Suzuki et al., 2006, Camacho et al., 2015). Amino acid
starvation and other stresses activate CsrB/C transcription via the stringent response
24
components ppGpp and DksA (Edwards et al., 2011). In addition, two DEAD-box RNA
helicases, DeaD and SrmB (Vakulskas et al., 2014), IHF (Zere et al., 2015) and cAMP-
Crp (Pannuri et al., 2016) activate or repress csrB/C transcription by distinct
mechanisms.
Regulation of the Turnover of CsrA Inhibitory sRNAs
The decay of CsrB/C RNAs has only been characterized in a few bacterial
species. In E. coli, CsrB/C turnover is mediated by RNase E and PNPase and facilitated
by protein CsrD (Suzuki et al., 2006). CsrD does not appear to be a nuclease but
renders CsrB/C susceptible to degradation by RNase E, thus affecting the expression of
CsrA-regulated genes in a predictable fashion (Suzuki et al., 2006). CsrD contains
GGDEF and EAL domains, which are often responsible for synthesis and degradation of
the secondary messenger cyclic dimeric (3′→5′) GMP (c-di-GMP) (Suzuki et al., 2006).
However, CsrD lacks the critical catalytic amino acid residues of GGDEF and EAL
domains and displays no c-di-GMP synthetic or hydrolytic activity (Suzuki et al., 2006).
Besides, its activity does not respond to c-di-GMP (Suzuki et al., 2006). So far, the
molecular mechanism of CsrD effects on CsrB/C degradation is unclear. CsrD binds
non-specifically to CsrB/C in vitro. Whether it has evolved from a typical GGDEF-EAL
domain protein to a protein with an alternative function, such as RNA binding, need to
be further studied. In addition, how CsrD activity is regulated is another open question.
CsrA weakly represses csrD expression in E. coli and Salmonella typhimurium (Suzuki
et al., 2006, Jonas et al., 2010) but does not seem to affect CsrB turnover in E. coli
(Suzuki et al., 2006). No other factors besides CsrA are known to affect CsrD activity.
CsrD orthologs are present in a number of bacterial families, including
Enterobacteriaceae, Vibrionaceae and Shewanellaceae (Suzuki et al., 2006). In E. coli,
25
the presence of CsrD drastically facilitates CsrB/C decay rates from half-lives of more
than 32 min to 1-4min during exponential phase growth. Nevertheless, in Pseudomonas
fluorescens, which lacks CsrD, these sRNAs are much more stable, with half-lives from
∼20 min to >60 min. Moreover, binding of the CsrA homolog RsmA stabilizes these
sRNAs by blocking RNase E cleavage in P. fluorescens (Duss et al., 2014, Reimmann
et al., 2005). However, CsrA displays on substantial effect on CsrB/C turnover in E. coli
(Suzuki et al., 2006). Whether this discrepancy is related to the presence of CsrD in E.
coli remains to be further explored.
CsrA Regulates its Own Expression
The level of CsrA in the cell is subject to a complex autoregulation. CsrA
represses its own translation by binding to the untranslated leader region of its mRNA
and blocking ribosome binding (Yakhnin et al., 2011). In the meanwhile, it activates its
own transcription indirectly through RpoS, which mediates the transcription of the P3
promoter of the csrA gene (Yakhnin et al., 2011). Furthermore, CsrA activity is
controlled by the negative feedback loop that exists within the Csr regulatory circuitry
(Gudapaty et al., 2001, Suzuki et al., 2002, Weilbacher et al., 2003). When CsrA activity
increases to certain level, it stimulates CsrB/C synthesis, therefore antagonizing its own
activity (Romeo et al., 2013). This complex autoregulation of its own expression and
activity fine-tunes the CsrA activity during different growth status and stress conditions.
Interaction of Csr System with Other Regulatory Pathways
CsrA participates in diverse regulatory pathways by directly binding to mRNAs of
the regulatory factors or components of these pathways (Romeo et al., 2013). A few
examples are described here. CsrA represses PGA-dependent biofilm formation at
26
multiple layers (Romeo et al., 2013). First, it directly binds to the untranslated leader of
pgaABCD mRNA and represses pgaA translation (Wang et al., 2005). In addition, it
inhibits the translation of NhaR, a positive regulator of pgaABCD transcription, by
occluding ribosome binding (Goller et al., 2006). Furthermore, CsrA reduces the
abundance of c-di-GMP, a secondary messenger that activates PGA synthesis and
biofilm formation, by controlling expression of genes responsible for c-di-GMP
metabolism (Jonas et al., 2008, Jonas et al., 2010). Besides, CsrA stimulates the Rho-
dependent transcription termination of pgaABCD mRNA by binding to the upstream
portion of its 5’-UTR and exposing the Rho recognization site within this region
(Figueroa-Bossi et al., 2014).
The same type of multiple layer regulation is also observed in CsrA regulation of
virulence formation in Legionella pneumophila (Sahr et al., 2017). CsrA regulates the
virulence formation by two routes, 1) by regulating the expression of several major
regulatory proteins (FleQ, LqsR, LetE and RpoS) related to virulence formation and 2)
by directly interacting with the transcripts of over 40 Dot/Icm type IV secreted effector
proteins and modulating their synthesis.
In addition, a reciprocal regulatory interaction is observed between the Csr
system and the stringent response (Edwards et al., 2011). During amino acid starvation
and other stresses, synthesis of the secondary messenger, (p)ppGpp, is stimulated and
modulates transcription of a large set of gene for stress resistant response (Cashel &
Gallant, 1969, Potrykus & Cashel, 2008). Synthesis of CsrB/C is strongly stimulated by
(p)ppGpp and the (p)ppGpp-responsive transcription regulator DksA. In the meanwhile,
27
CsrA binds the transcripts of RelA and SpotT that are involved in ppGpp synthesis and
hydrolysis, and DksA in order to modulate their expression (Edwards et al., 2011).
Project Rationale and Objectives
sRNAs are important factors in posttranscriptional regulation. Their levels
respond to diverse environmental stimuli and thereby properly modulate expression of
particular gene sets, adapting cellular metabolism and physiology to environmental
niches (Gottesman & Storz, 2011, Storz et al., 2011). Unlike most of the sRNAs that act
by base-pairing target mRNAs, E. coli CsrB/C act indirectly by binding to and titrating
the posttranscriptional regulatory protein CsrA (Romeo et al., 2013). Fluctuations of
CsrB/C levels play important roles in governing CsrA activity and many physiological
processes in response to different environmental stimuli. Biosynthesis of CsrB/C is
extensively controlled by multiple regulators that use distinct mechanisms under diverse
stress conditions (Chavez et al., 2010, Huang et al., 2008, Edwards et al., 2011, Suzuki
et al., 2002, Camacho et al., 2015, Vakulskas et al., 2014, Zere et al., 2015, Pannuri et
al., 2016). Interestingly, the decay rates of CsrB/C also play a role in determining their
steady state levels (Suzuki et al., 2006). CsrD protein is identified as a specific protein
that facilitates CsrB/C turnover (Suzuki et al., 2006). The overall goal of this study is to
explore the molecular mechanism of the action of CsrD and the physiological role of
CsrD in the CsrB/C turnover pathway.
The first objective of this study is to understand the physiological role of CsrD in
the CsrB/C decay pathway in E. coli. A recent study discovered the CsrD homolog of
Vibrio cholera as a binding partner of EIIAGlc, a component of the carbohydrate
phosphotransferase system (PTS), which serves in the uptake and phosphorylation of
glucose (Pickering et al., 2012). But no physiological role was assigned to this
28
interaction. Thus, this study is to illustrate whether EIIAGlc regulates the turnover of
CsrB/C by interacting with CsrD in E. coli and to characterize the molecular mechanism
and physiological consequences of this regulation.
The second objective of this study is to explore the molecular mechanism by
which CsrD facilitates CsrB/C turnover. Degradation of many sRNAs requires additional
factors in addition to RNases, such as Hfq (Basineni et al., 2009) and RapZ (Göpel et
al., 2013), to recruit RNases for rapid degradation. However, until now, the molecular
mechanism of CsrD action in CsrB/C turnover pathway is not yet well-characterized.
CsrD binds to CsrB/C in vitro, albeit non-specifically (Suzuki et al., 2006). It contains
GGDEF and EAL domains, which are typically involved in c-di-GMP synthesis and
hydrolysis, respectively (Hengge, 2009). However, CsrD displays no activity in c-di-
GMP metabolism (Suzuki et al., 2006). Whether CsrD acts directly by binding to CsrB or
indirectly through other factors needs to be investigated.
The third major objective of this study is to explore the regulatory role of CsrD in
a genome-wide scale. CsrD was previously observed to regulate expression of a
number of CsrA target genes due to its effects on CsrB/C levels and CsrA activity
(Suzuki et al., 2006). Thus, the open question is whether CsrD has a broader regulatory
role in addtition to affecting CsrB/C turnover, and whether it regulates gene expression
primarily through CsrB/C and CsrA or other unknown factor(s) or pathway(s). Epistasis
analysis together with RNA-seq will be conducted to resolve these questiones.
29
Figure 1-1. Outline of the Csr system in E. coli (Vakulskas et al., 2015). CsrA generally binds to conserved GGA motifs in the 5’-untranslated or early coding region of target mRNAs, leading to changes in translation initiation (as shown here), RNA stability, and/or transcription elongation. CsrA activity is controlled primarily by the steady state levels of CsrB and CsrC (CsrB is shown here), which contain many high affinity CsrA binding sites that sequester CsrA from interacting with its lower affinity mRNA regulatory targets. The levels of CsrB/C are regulated at the level of transcription and turnover. Ribosomes are depicted in blue. I have obtained the permission to use this figure from American Society for Microbiology.
Leng, Y., Vakulskas, C.A., Zere, T.R., Pickering, B.S., Watnick, P.I., Babitzke, P., and Romeo, T. (2016) Regulation of CsrB/C sRNA decay by EIIAGlc of the phosphoenolpyruvate: carbohydrate phosphotransferase system. Mol Microbiol 99: 627-639.
30
CHAPTER 2 REGULATION OF CsrB/C sRNA DECAY BY EIIAGlc
Introduction
The Csr/Rsm system is present in diverse eubacteria, where it globally regulates
metabolism, biofilm formation, motility, virulence, quorum sensing, and stress response
systems (Romeo, 1998, Babitzke & Romeo, 2007, Romeo et al., 2013, Vakulskas et al.,
2015). The RNA binding protein CsrA/RsmA of the Csr system regulates gene
expression by interacting with sequences in mRNA, thus altering translation, mRNA
stability, and/or transcript elongation. CsrA governs genes responsible for bacterial
lifestyle transitions, repressing processes that are triggered upon entry into the
stationary phase of growth and conferring stress resistance, while activating processes
such as glycolysis, which support vigorous growth. In E. coli, CsrA activity is mainly
controlled by the noncoding sRNAs, CsrB and CsrC, which contain multiple CsrA
binding sites, allowing them to antagonize CsrA activity by sequestering it away from
lower affinity mRNA targets (Liu et al., 1997, Weilbacher et al., 2003). Fluctuations in
CsrB/C levels play a central role in regulating Csr system and the bacterial lifestyle.
Multiple factors ensure appropriate expression of csrB/C. Transcription is
activated by the BarA-UvrY TCS in response to carboxylic acids (Suzuki et al., 2002,
Chavez et al., 2010, Martínez et al., 2014). In a complex negative feedback loop, CsrA
regulates csrB/C transcription by activating both the expression of the response
regulator UvrY and its phosphorylation by the sensor-kinase BarA (Suzuki et al., 2006,
Camacho et al., 2015, Suzuki et al., 2002). Amino acid starvation and other stresses
activate csrB/C transcription via the stringent response components ppGpp and DksA
31
(Edwards et al., 2011) and two DEAD-box RNA helicases, DeaD and SrmB, activate
csrB/C transcription by distinct mechanisms (Vakulskas et al., 2014).
In contrast to synthesis, the decay of CsrB/C RNAs is not well understood. A
specificity factor, CsrD, is necessary for degradation of CsrB/C by the housekeeping
nucleases RNase E and PNPase (Suzuki et al., 2006). CsrD does not appear to be a
nuclease, but renders CsrB/C susceptible to degradation by RNase E, thus affecting the
expression of CsrA-regulated genes in a predictable fashion. CsrD contains GGDEF
and EAL domains, which are often responsible for synthesis and degradation of the
secondary messenger cyclic dimeric (3’→5’) GMP (c-di-GMP). However, biochemical
and genetic studies indicated that CsrD displays no c-di-GMP synthetic or hydrolytic
activity and that CsrD activity is not regulated by c-di-GMP in vivo. At present, the
molecular mechanism of CsrD effects on CsrB/C degradation is unclear. CsrD bound
nonspecifically to CsrB/C in vitro (Suzuki et al., 2006). Accordingly, one hypothesis is
that CsrD evolved from the GGDEF-EAL domain family, becoming an RNA binding
protein. How CsrD activity is regulated is another open question. CsrA weakly represses
csrD expression in E. coli and Salmonella Typhimurium (Suzuki et al., 2006, Jonas et
al., 2010), but does not seem to affect CsrB turnover in E. coli (Suzuki et al., 2006), and
no other factors are known to affect CsrD activity.
Glucose is the preferred carbon and energy source for E. coli and is taken up
primarily by the glucose-specific phosphoenolpyruvate-dependent sugar-
phosphotransferase system, PTS (Deutscher et al., 2006, Deutscher et al., 2014,
Lengeler & Jahreis, 2009). This system consists of two cytoplasmic proteins, enzyme I
(EI) and histidine phosphocarrier protein (HPr) that are used for transporting many
32
sugars, and two glucose-specific proteins, enzyme IIAGlc (EIIAGlc) and the membrane-
bound enzyme IIBCGlc (EIIBCGlc). Glucose uptake is coupled to its phosphorylation. The
phosphoryl group is donated by PEP and transferred to glucose via a phosphorylation
cascade formed by EI, HPr, EIIAGlc, and EIIBCGlc proteins. Thus, the phosphorylation
state of the PTS proteins depends both on extracellular carbon availability and the
metabolic state of the cell.
The glucose-PTS proteins also mediate regulatory functions (Gabor et al., 2011,
Deutscher et al., 2014). EIIAGlc is a central regulator of carbon metabolism.
Unphosphorylated EIIAGlc mediates inducer exclusion by binding to and inhibiting
transporters of non-PTS sugars (Deutscher et al., 2014, Deutscher et al., 2006). It also
inhibits metabolism of alternative carbon sources, e.g. by binding to glycerol kinase
(Postma et al., 1984). In contrast, phosphorylated EIIAGlc (EIIAGlc-P) binds to and
stimulates the activity of adenylate cyclase, which produces cAMP. This compound acts
as a secondary messenger that binds to the cAMP receptor protein (Crp), forming a
transcription factor (cAMP-Crp) that exerts global effects on the proteome, ensuring
efficient resource utilization (Krin et al., 2002, Park et al., 2006, Bao & Duong, 2013).
With the discovery of novel EIIAGlc binding partners in various species, EIIAGlc has been
found to participate in chemotaxis (Neumann et al., 2012), respiration/fermentation (Koo
et al., 2004), biofilm formation (Pickering et al., 2012) and virulence (Kim et al., 2010,
Mazé et al., 2014). The EIIBCGlc protein also carries out a variety of regulatory functions
(Lux et al., 1995, Nam et al., 2001, Tanaka et al., 2000, Lee et al., 2000).
In a screen for EIIAGlc binding partners in Vibrio cholerae, a CsrD homologue,
MshH was identified (Pickering et al., 2012) . While no function was assigned to this
33
interaction, it was hypothesized that EIIAGlc might affect the decay of Csr sRNAs. Here,
we present the results of a detailed investigation of the role of EIIAGlc on CsrB/C decay
in E. coli. Unphosphorylated EIIAGlc binds specifically to the EAL domain of CsrD and
stimulates CsrB/C turnover. We propose that this mechanism helps to increase the
concentration of free CsrA when it is needed to support growth, and simultaneously
poises the Csr system for rapid response to changing environmental conditions.
Materials and Methods
Bacterial Strains and Culture Conditions
The bacterial strains used in this study are listed in Table B-1. Bacterial strains
were routinely grown in Luria-Bertani (LB) broth unless otherwise indicated. For
synthetic minimal medium, minimal medium A (Hogema et al., 1998) and M9 minimal
medium supplemented with indicated carbohydrates were used. When necessary, the
following antibiotics were added to the growth media: ampicillin (100 μg mL−1),
tetracycline (15 μg mL−1), kanamycin (50 μg mL−1), and chloramphenicol (25 μg mL−1).
E. coli and V. cholerae strains were grown at 37°C and 27°C, respectively. Stationary
phase cultures were routinely used to inoculate LB broth or minimal media unless
otherwise indicated. For strains carrying cyaA deletion or crp disruption, exponentially
growing cultures were used to inoculate LB broth supplemented with or without 10 mM
cAMP to minimize the growth defect.
Construction of Plasmids and Mutant Strains
The plasmids and related primers and restriction sites used in this study are
listed in Tables B-2 and B-3. The plasmid pCRR, used for complementation of a crr
deletion strain, carries the crr gene under a mutant lacUV5 promoter, in which -8 A of
the -10 hexamer consensus was replaced with -8 T for decreased promoter activity
34
(Moyle et al., 1991). To generate pCRR, the crr gene was amplified from chromosomal
DNA of E. coli MG1655 and ligated into vector pBR322. Plasmid pCRRH91A expressed
the mutant crr gene, producing the protein EIIAGlcH91A. This plasmid was constructed
similarly to pCRR except that the His91Ala (CAC to GCC) substitution was introduced
by the megaprimer PCR procedure (Ke & Madison, 1997). Plasmid pBYH4 used for
complementation of the csrD deletion strain was described previously (Suzuki et al.,
2006). To construct plasmid pETCRR for expression of C-terminal His-tagged EIIAGlc,
crr was amplified and cloned into plasmid pET24a. Plasmids overexpressing CsrD
variants were generated by amplifying the coding regions corresponding to CsrDΔTM
(residues 156-646), CsrDΔHAMP (residues 192-646), CsrDΔCoil (residues 156-199 and
220-656), CsrDΔGGDEF (residues 156-223 and 393-646), CsrDΔEAL (residues 156-385)
and EAL (residues 393-646) by standard PCR or overlapping PCR (Urban et al., 1997),
and cloning the resulting products into vector pmal-c5x, yielding N-terminally MBP-
tagged CsrD variants.
E. coli gene deletions were created by the standard P1vir transduction procedure
or the lambda Red system as described (Datsenko & Wanner, 2000). Chromosomal C-
terminal FLAG-tagged fusions were generated using the phage lambda Red system as
described (Datsenko & Wanner, 2000, Uzzau et al., 2001). The mutant strain H91A
carrying a chromosomal point mutation of crr, producing EIIAGlc H91A protein, was
constructed by using overlapping PCR mutagenesis and the pKOV gene replacement
protocol as described (Urban et al., 1997, Link et al., 1997).
35
V. cholerae ∆crr and ∆mshH mutants were created in the C6706str2 wild-type
background (Thelin & Taylor, 1996) by double homologous recombination as previously
described (Haugo & Watnick, 2002, Pickering et al., 2012).
Expression and Purification of EIIAGlc and CsrD Variants
EIIAGlc was overproduced in E. coli strain BL21 (DE3) grown in 1L of M9 minimal
medium supplemented with 0.8% glucose (w/v). Three hours after the induction with
1mM IPTG at OD600 ~ 0.6, cells were harvested and lysed using a French Press. After
centrifugation (20,000 × g, 30 min, 4 ̊C), the soluble fraction of the lysate was applied to
a HisTrap column (1 mL, GE Healthcare) and eluted with a gradient of imidazole (20-
500 mM). Eluted proteins were further purified by gel filtration chromatography
(Superdex 75 10/300, GE Healthcare), dialyzed against dialysis buffer (20 mM Tris-HCl,
pH7.5, 100 mM NaCl, 1 mM DTT, 10% glycerol) and stored for subsequent
experiments.
Overproduction of CsrD variants was from E. coli BL21 (DE3) strains containing
the corresponding plasmids, which were grown in LB medium supplemented with 0.2%
glucose (w/v). Cells were induced with 0.3 mM IPTG at OD600 ~ 0.6 and lysed as
mentioned above. The soluble fraction of the lysate was applied to an MBP Trap column
(1 mL, GE Healthcare) and eluted with a gradient of 0-10 mM maltose. To obtain
homogeneous proteins, eluted proteins were further purified by gel filtration
chromatography (Superdex 200 10/300, GE Healthcare) and dialyzed against dialysis
buffer.
Northern Blotting
Northern blot analysis was performed as previously described, with minor
modifications (Vakulskas et al., 2014). Bacterial culture were immediately stabilized by
36
the addition of RNA protect TM Bacteria Reagent (Qiagen) or 0.125 volumes of stop
solution (10% phenol/90% ethanol). Total RNA was isolated using the RNeasy mini kit
(Qiagen) according to the manufacturer’s instructions. Total cellular RNA (1-2 μg) was
separated on denaturing 5% acrylamide/7 M urea gels and transferred to a positively
charged nylon membrane (Roche Diagnostics) by electroblotting. The membrane was
cross-linked using UV light and hybridized overnight at 68°C (CsrB/C of E. coli) or 70°C
(CsrB/C/D of V. cholerae) using a DIG-labeled antisense RNA probe (Table B-3), which
was prepared with the DIG Northern Starter kit (Roche Diagnostics). Transcripts were
detected using the DIG Northern Starter kit (Roche Diagnostics) according to the
manufacturer’s instructions. Blots were imaged using the ChemiDoc XRS+ system (Bio-
Rad) and RNAs were quantified using Quantity One image analysis software (Bio-Rad).
Prior to hybridization, the rRNAs (16S and 23S) were stained by methylene blue, which
served as loading controls for signal correction.
Western Blotting
Western blot analysis was performed using standard laboratory protocols as
described (Vakulskas et al., 2014). Briefly, total cellular proteins were separated by
SDS-PAGE and transferred to 0.2 μm polyvinylidene difluoride membranes (Bio-Rad)
by electroblotting. Blots for FLAG epitope-tagged proteins used the anti-FLAG M2
monoclonal antibody (Sigma) at 1:2,000 dilution. Blots for RpoB used anti-RpoB
monoclonal antibodies (Neoclone) at 1:50,000 dilution. Western blots were detected
using horseradish peroxidase-linked secondary antibodies and the Super Signal West
Femto Chemiluminescent Substrate (Pierce) as recommended by the manufacturer.
Proteins were quantified using Quantity One image analysis software (Bio-Rad).
37
Protein Pull-down Assays
Interaction between CsrD variants and EIIAGlc was assessed using purified His-
tagged EIIAGlc protein to pull down MBP-tagged CsrD variants. In a 60 μL reaction, 8
μM EIIAGlc and 4 μM CsrD variants were incubated with 15 μL Ni-NTA resin (Qiagen) in
binding buffer (50 mM MES pH 6.5, 1 mM DTT, 20 mM Imidazole) at 4°C for 1hour.
Unbound proteins remaining in the supernatant solution were collected after brief
centrifugation of the resin. The resin was washed three times with 1 mL washing buffer
(50 mM MES pH 6.5, 1mM DTT, 60 mM Imidazole), and the bound proteins were eluted
with 45 μL of elution buffer (25 mM Tris-HCl pH 8.0, 500 mM NaCl, and 500 mM
Imidazole).
To determine the phosphorylation dependence of the interaction between EIIAGlc
and CsrD, the MBP-tagged CsrDΔTM was used to pull down His-tagged EIIAGlc. His-
tagged EIIAGlc (8 μM) was incubated in reactions containing EI (1 μM), Hpr (8 μM) and
either 5 mM PEP or 5 mM pyruvate in 60 μL buffer (20mM Tris-HCl pH8.0, 2 mM MgCl2,
1 mM DTT) at room temperature for 10 min. These reactions were designed to produce
the phosphorylated or unphosphorylated form of EIIAGlc, respectively. Reactions were
dialyzed against the binding buffer (50mM MES pH6.5, 1mM DTT) with Slide-A-Lyzer
dialysis cassette (Thermo) for 1 hour and subsequently incubated at 4°C for 1 hr with
CsrDΔTM (95 μg) pre-bound to amylose resin. Unbound proteins were collected after
brief centrifugation of the resin and bound proteins were eluted with 60 μL of elution
buffer (20 mM Tris-HCl pH7.5, 200 mM NaCl, 10 mM maltose, 1 mM DTT) after
extensive rinsing of the resin with binding buffer. Unbound and bound proteins in the
pull-down reactions were detected by SDS-PAGE (10% or 15%, as required) and
38
Coomassie blue staining. Proteins were quantified using Quantity One image analysis
software (Bio-Rad).
Determination of the EIIAGlc Phosphorylation State
The phosphorylation state of EIIAGlc was determined as previously described
(Hogema et al., 1998), with modifications. Briefly, 0.2 mL of bacterial culture (OD600 ~
0.5) was treated with the addition of 20 μL of 10 M NaOH followed by 1 mL of ethanol
and 180 μL of 3 M sodium acetate, pH 5.2. After chilling at -80°C for 2 hr, precipitates
were collected by centrifugation, rinsed with 70% ethanol, and suspended in 100 μL of
sample buffer (0.16 M Tris HCl, pH 7.5, 4% SDS, 20% glycerol, and 10% 2-
mercaptoethanol). To achieve a good separation of the two forms of EIIAGlc, samples
were fractionated on SDS-PAGE gels containing 50 μM of Phos-tag reagent as
previously described (Vakulskas et al., 2014). Subsequently, gels were washed with
Western transfer buffer (25 mM Tris, 192 mM Glycine, 20% methanol, and 0.1% SDS)
containing 1 mM EDTA for 10 min, followed by a second wash with transfer buffer for 20
min. Western blot analysis was then performed as described above.
Gel Filtration Analysis of EIIAGlc in Complex with the EAL Domain or Other CsrD Variants
As discussed below, a site-directed crr mutant allele, encoding an EIIAGlc protein
that cannot be phosphorylated, H91A, also complemented the crr deletion (Fig. 2-1A-D).
For analysis of the EIIAGlc-EAL complex, a 0.5 mL reaction containing 33 μM EAL, 78
μM EIIAGlc or both proteins were incubated at 4 °C for 30 min and subjected to gel
filtration analysis using an AKTA-FPLC system (Superdex 200, HiLoadTM 16/60, 120
mL, GE Healthcare), and subsequently eluted at 4°C with a flow rate of 1 mL/min in
buffer containing 20 mM Tris HCl pH 7.5, 100 mM NaCl, 1mM DTT. For CsrD variants,
39
0.5 mL samples containing purified CsrDΔTM, CsrDΔEAL, CsrDΔHAMP, CsrDΔCoil, or
CsrDΔGGDEF was separated on the same system. Fractions (3 mL) were collected and
analyzed by SDS-PAGE and Coomassie blue staining. For EIIAGlc-EAL analysis, the
column was pre-calibrated using 5 gel filtration molecular weight markers (1, sweet
potato β-amylase, 200 kDa; 2, yeast alcohol dehydrogenase, 150 kDa; 3, bovine serum
albumin, 66 kDa; 4, carbonic anhydrase from bovine erythrocytes, 29 kDa; 5, horse
heart cytochrome C, 12.4 kDa), and blue dextran 2000. For CsrD variants analysis, the
column was calibrated using 5 molecular weight markers (1, equine spleen apoferritin,
443 kDa; 2, sweet potato β-Amylase, 200 kDa; 3, alcohol yeast dehydrogenase, 150
kDa; 4, bovine serum albumin, 66 kDa; 5, carbonic anhydrase from bovine erythrocytes,
29 kDa), and blue dextran 2000. The relative molecular masses of proteins or protein
complexes were calculated by logarithmic interpolation from the standards.
Results
EIIAGlc Activates CsrB/C Decay via CsrD
Because the role of CsrD in CsrB/C decay has been investigated only in E. coli,
we decided to determine if EIIAGlc participates in the degradation of CsrB/C in this
species. To do so, we first determined the stability of CsrB/C in the presence or
absence of EIIAGlc (Δcrr) after the addition of rifampicin to the exponentially growing
cultures. Deletion of crr decreased CsrB and CsrC decay rates by about 5-fold and 3-
fold, respectively (Fig. 2-1A-D). Ectopic expression of crr complemented the crr defect,
confirming that EIIAGlc somehow regulates CsrB/C decay (Fig. 2-1A-D). As discussed
below, a site-directed crr mutant allele, encoding an EIIAGlc protein that cannot be
phosphorylated, H91A, also complemented the crr deletion (Fig. 2-1A-D).
40
As shown in Fig. 2-1E, while EIIAGlc greatly stimulated CsrB/C decay, it modestly
reduced the levels of CsrB/C in the cell. A similar observation was also made previously
in a csrD mutant strain, and was shown to be the result of the Csr regulatory circuitry
(Suzuki et al., 2006). CsrA indirectly activates transcription of CsrB/C via the BarA-UvrY
TCS (Suzuki et al., 2002). Thus, when CsrB/C decay is inhibited, these sRNAs
accumulate and sequester CsrA, causing a decrease in their transcription and an
attenuated effect on their levels.
We next performed an epistasis experiment to determine whether the effect of
EIIAGlc on CsrB/C decay was dependent on CsrD. A Δcrr ΔcsrD double deletion strain
was severely defective in CsrB/C decay (Fig. 2-2A-B), as reported previously for the
ΔcsrD mutant (Suzuki et al., 2006). While ectopic expression of crr in the Δcrr ΔcsrD
strain failed to restore CsrB/C decay, ectopic overexpression of csrD enhanced CsrB/C
decay rates to wild-type levels (Fig. 2-2A and B). This finding suggested that CsrD
functions downstream of EIIAGlc in CsrB/C turnover.
A possible explanation for the epistasis results is that EIIAGlc affects CsrD levels
in the cell by altering its stability or synthesis. However, deletion of crr had no effect on
the levels of a chromosomally encoded and biologically functional CsrD-FLAG protein
(Fig. 2-2C and Fig. A-1).
In vitro Binding of EIIAGlc to CsrD Requires the EAL Domain
Because EIIAGlc regulates the activities of several proteins via direct binding, it
was reasonable to speculate that EIIAGlc affects CsrB/C decay via direct binding to CsrD
in E. coli. To test this idea, we examined the binding of EIIAGlc and CsrD in an in vitro
binding assay or pull-down assay using His-tagged EIIAGlc and a soluble recombinant
CsrD protein (CsrDΔTM) in which the N-terminal transmembrane domains of CsrD were
41
replaced with a maltose binding protein (MBP) tag (Fig. 2-3A). Previous studies showed
that the transmembrane domains of CsrD were dispensable for its activity when the
protein was ectopically expressed (Suzuki et al., 2006). In this assay, CsrD was mixed
with Ni-NTA resin with or without His-tagged EIIAGlc. As shown in Fig. 2-3B, the CsrDΔTM
variant was retained by the Ni-NTA resin when EIIAGlc was bound to it, but remained in
the unbound fraction when EIIAGlc was absent, indicating that EIIAGlc bound directly to
CsrD.
To determine which domain of CsrD protein is involved in the interaction with
EIIAGlc, we tested similar MBP fusions of CsrD lacking the EAL (CsrDΔEAL), GGDEF
(CsrDΔGGDEF), or HAMP-like domain (CsrDΔHAMP and CsrDΔCoil), using the pull-down
assays. While the other CsrD variants retained the ability to bind to EIIAGlc, CsrDΔEAL
lost all detectable binding, suggesting that the EAL domain is involved in this interaction
(Fig. 2-3B-C). Moreover, the EAL domain alone bound to EIIAGlc in this assay (Fig. 2-
3D). These results indicated that EIIAGlc binds specifically to the EAL domain of CsrD.
To examine the binding reaction of EIIAGlc with CsrD in more detail, the size and
composition of the EIIAGlc-EAL complex was analyzed by gel-filtration chromatography.
The free EAL and EIIAGlc eluted at positions corresponding to sizes of their monomeric
forms (EAL, 72kDa; EIIAGlc, 20 kDa) (Fig. 2-3E and F). When EIIAGlc and EAL were
mixed to allow binding, and fractionated on Superdex 200 (HiLoadTM 16/60, GE
Healthcare), a new peak was observed at a position corresponding to a size of 98kDa,
approximately that of a heterodimer of EIIAGlc-EAL (92kDa). To determine the ratio of
EIIAGlc and EAL in the complex, column fractions corresponding to the presumptive
EIIAGlc-EAL complex and the free EIIAGlc were analyzed by SDS-PAGE with
42
Commassie blue staining and quantification of the stained proteins (Fig. 2-3F). This
experiment revealed the molar ratio of EAL bound to EIIAGlc in peak fractions (11 and
12) to be 1:1 suggesting that EIIAGlc binds to the EAL domain of CsrD in a one to one
ratio.
In the pull down assay, the relative amount of CsrDΔTM or CsrDΔGGDEF bound by
EIIAGlc was much greater than that of proteins that lacked the intact HAMP-like domain,
CsrDΔHAMP or CsrDΔCoil (Fig. 2-3C). The HAMP domain typically promotes dimerization
or protein-protein interactions and plays important roles in signal transduction (Hulko et
al., 2006). We used gel filtration assays to determine the in vitro oligomeric states of
CsrD variants containing or lacking the HAMP-like domain. All CsrD variants containing
the HAMP-like domain, CsrDΔTM, CsrDΔGGDEF and CsrDΔEAL, eluted at volumes
consistent with their tetrameric forms (Fig. 2-4 and Table 2-1). In contrast, the CsrD
variants with a disrupted HAMP-like domain, CsrDΔHAMP and CsrDΔCoil, eluted as
apparent monomers (Fig. 2-4 and Table S4). The precise way in which tetramerization
affected EIIAGlc binding by the CsrD variants was not further investigated.
EIIAGlc Regulates CsrB/C Decay in a Phosphorylation-dependent Manner
EIIAGlc typically modulates the activity of its binding partners in a
phosphorylation-dependent manner. Accordingly, we tested the effect of the
phosphorylation state of EIIAGlc on CsrB/C decay. We first investigated the impact of the
unphosphorylated EIIAGlc on CsrB/C turnover using a site-directed mutant protein that
could not be phosphorylated (EIIAGlc H91A). Plasmid complementation of the Δcrr strain
with EIIAGlc H91A restored CsrB/C decay rates to slightly higher than in the wild-type
strain (Fig. 2-1A-D), demonstrating that phosphorylation of EIIAGlc was dispensable for
activation of CsrB/C turnover.
43
Next, we performed pull-down assays to determine the effect of EIIAGlc
phosphorylation on binding to CsrD. In these experiments, CsrD containing an N-
terminal MBP tag was bound to amylose resin and then mixed with EIIAGlc in reactions
that were designed to produce either the phosphorylated or unphosphorylated form of
this protein. In one reaction, E. coli Hpr, EI and PEP were mixed to provide
phosphorylated EIIAGlc. In the other reaction, pyruvate was added instead of PEP to
maintain EIIAGlc in the unphosphorylated form. Strikingly, while most of the
unphosphorylated EIIAGlc bound to CsrD, no binding was observed between the
phosphorylated EIIAGlc and CsrD (Fig. 2-5). These results were in agreement with the
observation that EIIAGlc did not require phosphorylation for activation of CsrB/C turnover
in vivo (Figs. 2-1A-D), and indicated that the binding of unphosphorylated EIIAGlc to
CsrD activates CsrB/C sRNA decay.
cAMP-Crp Modestly Represses CsrB Turnover
While the unphosphorylated EIIAGlc bound to CsrD in vitro and was able to
activate CsrB/C decay in vivo, we wondered if the phosphorylated form of EIIAGlc might
affect CsrB/C turnover via its important role in cAMP-Crp production (Krin et al., 2002,
Park et al., 2006). Deletion of cyaA or crp modestly increased the CsrB decay rate by 2-
fold (Fig. 2-6A), while exhibiting weak or negligible effects on CsrC decay (Fig. 2-6B).
The increased decay rate of CsrB in the cyaA mutant was restored by exogenous cAMP
(10 mM), confirming that cAMP-Crp somehow inhibits CsrB decay. Deletion of cyaA or
crp in the Δcrr background had twofold effects on CsrB decay rates that were similar to
those in the wild-type background, and deletion of crr had similar fivefold effects on
CsrB decay in both the wild-type strain and its isogenic crp and cyaA mutants (Fig. 2-
6A). These findings confirmed that the major effect of EIIAGlc on CsrB decay is mediated
44
independently of cAMP-Crp. The modest effect of cAMP-Crp in the Δcrr background
was likely due to basal adenylate cyclase activity in the crr mutant (Lévy et al., 1990,
Feucht & Saier, 1980, Reddy & Kamireddi, 1998).
CsrB/C Decay is Regulated in Response to Carbon Availability via the Phosphorylation State of EIIAGlc
The phosphorylation state of EIIAGlc is determined by carbon sources that are
taken up and metabolized (Hogema et al., 1998, Deutscher et al., 2014). Preferred
carbon sources such as glucose lead to net dephosphorylation of PTS proteins,
including EIIAGlc, whereas unfavorable carbon sources or carbon starvation conditions
cause the accumulation of phosphorylated EIIAGlc. Because the unphosphorylated
EIIAGlc bound to CsrD in vitro and promoted CsrB/C decay in vivo (Figs. 2-1 and 2-5),
we expected that CsrB/C decay rates should be elevated in the presence of glucose.
Consequently, we first examined CsrB/C decay in minimal medium supplemented with
0.2% glucose, glycerol or succinate. Both the phosphorylation state of EIIAGlc and
CsrB/C decay rates responded predictably to these carbon sources; more rapid decay
was observed in glucose compared to glycerol or succinate (Fig. 2-7).
To verify that the phosphorylation state of EIIAGlc determines the decay rates of
CsrB/C in response to different carbon sources, we examined CsrB/C turnover in a
strain that expresses the mutant EIIAGlc protein, H91A, which cannot be phosphorylated.
Because the strain expressing H91A has a significant growth defect in minimal medium
(data not shown) the WT and H91A strains were first grown in LB broth to exponential
phase and then washed and inoculated into minimal medium lacking a carbon
compound or containing 0.2% glucose or succinate. The phosphorylation state of EIIAGlc
was determined from growth in LB (Fig. 2-8A) and 10 min after inoculation into minimal
45
media (Fig. 2-8B). EIIAGlc phosphorylation in LB was ~40% and increased to ~90% in
media with succinate or lacking a carbon source, while it decreased to 4% at 10 min
after glucose exposure. Decay rates of CsrB/C were determined 10 min after
inoculation (Fig. 2-9). The decay rates of CsrB and CsrC in the wild-type strain (WT)
were ~3.5 and 2.5-fold greater, respectively, in medium with glucose vs. no carbon
source. A more modest, but reproducible difference (~2 fold) was observed for CsrB/C
decay rates in glucose compared to succinate. These data support the observations
described above, showing that CsrB/C decay rates vary in response to different carbon
conditions, although the difference in decay between succinate and carbon-deficient
media does not seem to be explained by EIIAGlc phosphorylation alone (Fig. 2-9), as
both conditions resulted in similar EIIAGlc-P levels (Fig. 2-8B). Most importantly, CsrB/C
decay in the H91A strain was rapid and virtually identical in all three media, confirming
that the phosphorylation state of EIIAGlc determines CsrB/C decay in response to carbon
substrate availability. The levels of CsrB/C RNAs (Fig. 2-9E) were consistent with these
decay rates, but as observed previously (Fig. 2-1), the effects of turnover may be
attenuated via the feedback loop of the Csr circuitry (Fig. 2-1F) (Suzuki et al., 2002,
Suzuki et al., 2006).
EIIAGlc and MshH Promote Csr sRNA Decay in Vibrio cholerae
EIIAGlc, RNase E and CsrD orthologs are widespread in Enterobacteriaceae,
Vibrionaceae, and Shewanellaceae species (Suzuki et al., 2006, Vakulskas et al.,
2015), suggesting that a common mechanism may exist for Csr sRNA decay in
members of these bacterial families. As a proof of principle, we tested the effects of
EIIAGlc and the CsrD homolog, MshH, on decay of the V. cholerae sRNAs, CsrB, CsrC
and CsrD (Lenz et al., 2005). The V. cholerae sRNAs exhibited longer half-lives
46
compared to the E. coli sRNAs under our growth conditions (Fig. 2-10). Nevertheless,
deletion of mshH, crr or both genes greatly decreased CsrB and CsrD turnover. These
effects were not apparent for CsrC, which was already extremely stable in the wild-type
strain. This experiment demonstrated the potential of EIIAGlc and CsrD to activate the
decay of Csr sRNAs in this important member of the Vibrionaceae family.
Discussion
Here, we identified a new regulatory function for EIIAGlc, in which binding to the
sRNA decay protein CsrD stimulates CsrB/C decay when glucose is present. This
mechanism should enhance the concentration of free CsrA, which activates glycolysis
and represses gluconeogenesis, secondary metabolism, and stress resistance
responses such as biofilm formation (Babitzke & Romeo, 2007, Vakulskas et al., 2015,
Romeo et al., 2013). Because CsrA regulates lifestyle transitions in many bacterial
species and interacts with hundreds of transcripts in E. coli (Babitzke & Romeo, 2007,
Patterson-Fortin et al., 2013, Edwards et al., 2011, Vakulskas et al., 2015), we propose
that this represents a particularly important role for EIIAGlc. A high rate of turnover can
facilitate rapid changes in transcript levels. Therefore, EIIAGlc-CsrD interactions should
not only allow CsrB/C decay rates to be reset in response to changing glucose
availability, but may poise the Csr system for rapid responses to other cues or
conditions when glucose is present. The V. cholerae CsrD ortholog, MshH and EIIAGlc
also activated the decay of CsrB and CsrD sRNAs of (Fig. 2-10). We suspect that the
mechanism described for E. coli CsrB/C turnover operates in many species of
Enterobacteriaceae, Vibrionaceae and Shewanellaceae.
The conclusion that only the unphosphorylated form of EIIAGlc is able to promote
CsrB/C decay through binding interactions with CsrD was based on a combination of
47
biochemical and genetic evidence. CsrD bound only to the unphosphorylated form of
EIIAGlc in vitro (Fig. 2-5). Furthermore, a non-phosphorylatable protein, EIIAGlc-H91A,
sustained CsrB/C decay rates that were similar to or even greater than the wild-type
protein (Figs.2-1A-D). The presence of glucose also caused net dephosphorylation of
EIIAGlc and supported rapid decay of CsrB/C sRNAs relative to starvation conditions or
alternative carbon sources (Figs. 2-7-9). Importantly, EIIAGlc-H91A supported high
decay rates under all of these conditions, confirming that the effects of carbon
availability on CsrB/C decay are mediated thorough altered phosphorylation of EIIAGlc
(Fig. 2-9). A previous study with MshH of V. cholera concluded that both the
phosphorylated and unphosphorylated forms of EIIAGlc bind to the CsrD homolog. This
conclusion was based on the observation that in a two-hybrid assay, MshH interacted
with mutant proteins designed to mimic the unphosphorylated (EIIAGlc-H91A) or the
phosphorylated (EIIAGlc-H91D) forms of EIIAGlc (Pickering et al., 2012). However, we
caution that EIIAGlc-H91D does not appear to mimic EIIAGlc-P. Another putative EIIAGlc-P
mimic, EIIAGlc-H91E, was unable to activate adenylate cyclase (Reddy & Kamireddi,
1998). Similarly, effects of EIIANtr-P were not mimicked by replacing its
phosphorylatable His residue with either Asp or Glu (Lüttmann et al., 2009). Finally, we
constructed EIIAGlc-H91D in E. coli and found that it behaved similarly to
unphosphorylated EIIAGlc rather than EIIAGlc-P in CsrB/C decay (Fig. A-2).
The phosphorylated form of EIIAGlc activates cAMP synthesis by binding to
adenylate cyclase. Because cAMP and Crp modestly repressed CsrB decay (Fig. 2-6),
we propose that EIIAGlc-P indirectly and modestly represses CsrB turnover, reinforcing
the positive effect of unphosphorylated EIIAGlc on CsrB decay. Because a potential Crp
48
binding site was predicted in the untranslated leader region of csrD (data not shown),
we tested the possibility that Crp inhibits CsrB decay by controlling csrD expression.
Weakly positive to negligible effects of Crp were observed on CsrD levels (Fig. A-3),
which might contribute to the effect of Crp on CsrB decay.
Given that EIIAGlc acts via CsrD without altering its levels in the cell and that
overexpression of csrD restored CsrB/C decay in a strain deleted for the EIIAGlc gene,
crr (Fig. 2-2), we propose that EIIAGlc functions as an allosteric activator of CsrD,
perhaps similar to the role of EIIAGlc-P in activating adenylate cyclase (Saier, 1989, Park
et al., 2006). Structural studies of EIIAGlc show that it possesses a concave face that
allows it to interact with globular target proteins (Hurley et al., 1993, Chen et al., 2013,
Wang et al., 2000, Cai et al., 2003). Other EIIAs seem unable to duplicate this function
(Deutscher et al., 2006). We deleted the genes for five other EIIAs (fruB, mtlA, chbA,
manX and ptsN) and found that none of them regulated CsrB/C decay (data not shown).
This study also expands our understanding of the functionality of the EAL
domain. This is not the first report of a catalytically inactive EAL domain performing a
regulatory role via protein-protein interactions (Römling et al., 2013). The enzymatically
inactive EAL domain protein YdiV of E. coli binds to the transcription activator FlhD4C2
and prevents it from binding to target DNA (Li et al., 2012). Similarly, the EAL domain of
the E. coli photoreceptor YcgF (BluF) binds to the MerR-like repressor YcgE (BluE) in
the presence of blue light and prevents it from binding to DNA (Tschowri et al., 2009).
Complex regulation exists for the EAL domain of FimX, which binds to c-di-GMP as well
as the PilZ protein, and is required for biogenesis of the Type IV pilus (Guzzo et al.,
2009). In all of these cases, the EAL domain-containing protein acts as a sensor that
49
uses its EAL domain to transmit information to another protein. In contrast, the EAL
domain of CsrD acts as a receiver, to detect signaling information from a sensory
protein. The modular structure of CsrD suggests that the effect of EIIAGlc might be
transmitted through other CsrD domains, such as the GGDEF domain, which is also
necessary for CsrD activity (Suzuki et al., 2006), although this possibility has not been
explored.
While the circuitry surrounding the Csr system is extensive, its role in carbon and
energy pathways is particularly wide-ranging and important (Edwards et al., 2011,
Patterson-Fortin et al., 2013, Yang et al., 1996, Romeo, 1996, Romeo et al., 2013,
Sabnis et al., 1995, Pernestig et al., 2003, Yang et al., 1996). Previous studies have
shown that carboxylic acid-containing end products of carbon metabolism, such as
acetate and formate, stimulate CsrB transcription via the BarA-UvrY TCS (Chavez et al.,
2010). Thus, the synthesis pathway and the newly discovered turnover pathway for
CsrB/C should mediate reinforcing positive effects on the levels of these sRNAs when
preferred carbon resources have been expended and end products have accumulated.
The resulting decrease in CsrA activity under this condition should promote the
transition from glycolytic metabolism and active growth to gluconeogenesis, glycogen
biosynthesis and the formation of a stress resistant phenotype. We caution that other
regulators influence the expression of CsrB/C RNAs, e.g. ppGpp, DksA, DeaD and
SrmB helicases, and impact the workings of this circuitry. An understanding of the
combinatorial effects of all of these factors, and perhaps unknown ones, will require
additional investigation.
50
Figure 2-1. EIIAGlc affects CsrB/C decay rates and levels in E. coli. A and C) Decay rates of CsrB/C were determined by Northern blotting of RNA extracted from exponential phase cultures (OD600 ~0.5) at various times following the addition of rifampicin. Culture identities: E. coli MG1655 (WT) or its Δcrr mutant with or without plasmid pBR322, pCRR or pCRRH91A (nonphosphorylatable EIIAGlc). The RNA half-lives were determined from the linear portions of their decay curves, shown in B and D. Standard deviations of values from two independent experiments are shown. E) CsrB/C steady state levels determined by Northern blotting of RNA from exponentially growing cultures (OD600 ~0.5), as above. F) A model for the Csr regulatory circuitry that includes EIIAGlc activation of CsrD-mediated CsrB/C decay. A broken line indicates an undefined mechanism(s).
51
Figure 2-2. EIIAGlc (crr) activates CsrB/C decay via CsrD, but does not enhance cellular
CsrD levels. A and B) Decay rates of CsrB (A) and CsrC (B) were determined by Northern blotting of RNA from E. coli MG1655 (WT), ΔcsrD, Δcrr ΔcsrD strains with or without plasmid pBR322, pCRR or pBYH4 (expressing CsrD), as described in Figure 2-1. C) Western blots depicting the effect of crr deletion on the level of CsrD protein. RpoB was used as loading control. CsrD protein levels in Δcrr relative to those in the wild-type strain (WT) are shown at the bottom. Standard derivations from triplicate experiments are indicated.
52
Figure 2-3. EIIAGlc interacts specifically with the EAL domain of CsrD. A) CsrD variants
used to identify the domain that binds to EIIAGlc. B-D) In vitro assays for binding of EIIAGlc to CsrD variants depicted in panel A. Each reaction contained 8 μM of CsrD variants and 16 μM of EIIAGlc. U and B: unbound and bound proteins. Control reactions were performed in the absence of EIIAGlc to confirm that CsrD varients do not bind nonspecifically to Ni-NTA resin. E) Gel filtration assay of EIIAGlc, EAL, and EIIAGlc-EAL mixture. Proteins were fractionated on a Superdex 200 column (HiLoadTM 16/60, 120 mL). The solid line corresponds to EAL domain alone; the dashed line corresponds to the EIIAGlc-EAL mixture. The chromatogram for EIIAGlc was not shown because this protein displays little absorbance at 280 nm. Arrows indicate elution volumes of molecular weight markers used to calibrate the column (Experimental procedures). F) SDS-PAGE and Coomassie blue staining of proteins from gel filtration chromatography fractions of the EIIAGlc-EAL mixture shown in panel E. Fractions (3 mL) were collected starting at 40 mL.
A B
C D
GGDEF EAL HAMP-like
Transmembrane
Coiled coil
CsrD
CsrDΔTM
CsrDΔEAL
CsrDΔGGDEF
CsrDΔHAMP
CsrDΔCoil
EAL
Binding to
EIIAGlc
ND
+
-
+
+
+
+
U B U B U B U B U B U B U B
+ _ + _ CsrDΔTM CsrDΔEAL
+ _ +
CsrDΔGGDEF
EIIAGlc
M (kDa)
CsrD variants:
EIIAGlc:
250 150 100 75
50
37
25 20
15
_
CsrDΔHAMP CsrDΔCoil
EIIAGlc
U B U B U B U B U B U B U B
+ _ + _ CsrDΔTM
+ _ + M (kDa)
CsrD variants:
EIIAGlc:
250 150 100 75
50
37
25 20
15
_
EIIAGlc
EAL
U B U B U B
+ M (kDa)
EAL:
EIIAGlc: _ +
_ +
250 150 100 75
50
37
25
20
15
53
Figure 2-3. Continued.
Figure 2-4. Gel filtration chromatography of CsrD variants. Each CsrD variant (1 mg)
was passed through a Superdex 200 column (HiLoadTM 16/60, 120 mL). Arrows indicate elution volumes of molecular weight markers used to calibrate the column (Materials and Methods).
E F 10 11 12 13 14 15 16 17 18
EIIAGlc
EAL 75
50
37
25 20
M (kDa)
15
54
Table 2-1. Molecular weight of CsrD in solution
Protein Molecular Mass (kDa) Quaternary structurec Calculateda Experimentalb
CsrDΔTM 99.3 359 Tetramer CsrDΔHAMP 95.1 107 Monomer CsrDΔCoil 97.0 107 Monomer CsrDΔGGDEF 80.3 344 Tetramer CsrDΔEAL 69.4 296 Tetramer
a. Molecular mass of the protein calculated from the primary sequence. b. Molecular mass determined using size exclusion chromatography. c. Deduced quaternary structure.
Figure 2-5. CsrD binds only to unphosphorylated EIIAGlc in pull-down assays. MBP-
tagged CsrD protein was bound to amylose resin and then mixed with EIIAGlc
in the nonphosphorylated (N) or phosphorylated state (P) to permit binding reactions to occur. The reactions were processed as described in the Experimental procedures. Control reactions were performed without CsrD to test for nonspecific binding of the two forms of EIIAGlc to amylose resin. Note that the two forms of EIIAGlc were resolved from each other on 15% SDS-PAGE gel. U and B refer to proteins that were unbound vs. bound by the amylose resin.
EIIAGlc:
_ + CsrDΔTM:
CsrD
EIIAGlc-P
EIIAGlc
EI
M (kDa) U B U B U B U B
N P N P
_ +
100 75 50
37
25
20
55
Figure 2-6. Effects of cAMP-Crp on CsrB/C decay. A and B) Decay rates of CsrB/C
were determined by Northern blotting of RNA from E. coli MG1655 (WT) and isogenic deletion mutants: cyaA, crp, cyaA, crr crp, crr cyaA, crr with or without added 10 mM cAMP, as described in Figure 2-1.
56
Figure 2-7. Effects of carbon sources on EIIAGlc phosphorylation and CsrB/C decay in
minimal media. A) Western blot depicting the phosphorylation state of EIIAGlc of E. coli MG1655 growing in minimal medium A supplemented with 0.2% glucose, glycerol or succinate. Exponential phase extracts were fractionated in gels with Phos-tag™ reagent and analyzed by Western blot. Relative level of total EIIAGlc was analyzed by western blot without using Phos-tag™ reagent. The percentage of phosphorylated (EIIAGlc-P) and unphosphorylated EIIAGlc (EIIAGlc) and relative total EIIAGlc protein levels are given. B and C) Northern blot depicting CsrB/C decay rates in E. coli MG1655 exponentially growing in minimal medium A supplemented with 0.2% glucose, glycerol or succinate. The RNA half-lives were determined as shown in Figure 2-1.
57
Figure 2-8. The phosphorylation state of EIIAGlc before and 10 min after shift from LB
broth into minimal media. A and B) Western blot depicting the phosphorylation state and relative level of EIIAGlc in E. coli MG1655 (WT), Δcrr and H91A (mutant strain that carrying a chromosomal EIIAGlcH91A point mutation). Extracts were prepared from cultures grown in LB broth (A) and at 10 min after reinoculation into minimal medium without carbon, with 0.2% glucose or succinate (B). The percentage of phosphorylated EIIAGlc relative to total EIIAGlc was determined as described in Figure 2-7.
58
Figure 2-9. CsrB/C decay rates and levels after shift from LB to minimal media. A-D)
Decay rates of CsrB (A and B) and CsrC (C and D) determined by Northern blotting of RNA from E. coli MG1655 (WT) and mutant strain H91A (unphosphorylatable EIIAGlc). Strains were grown in LB broth to exponential phase, washed and reinoculated into M9 minimal medium without carbon or with 0.2% glucose or succinate. Rifampicin was added 10 min after inoculation into minimal media and RNA half-lives determined as in Figure 2-1. E) CsrB/C steady state levels determined by Northern blotting of RNA from E. coli MG1655 (WT) in LB broth at exponential growth phase and 10 min after inoculation into minimal media.
A B
C D
E
Half-life (min)
9.4 ± 1.9
2.3 ± 0.1
5.3 ± 1.7
CsrB-WT
No C
Glc
Suc
0 2 4 6 8 16 32
Time (min)
3.2 ± 0.4
2.6 ± 0.1
2.8 ± 0.2
CsrB-H91A Half-life
(min) 0 2 4 6 8 16 32
No C
Glc
Suc
Time (min)
Half-life (min)
13.2 ± 2.0
4.7 ± 0.3
7.7 ± 2.0
CsrC-WT
No C
Glc
Suc
0 2 4 6 8 16 32
Time (min)
5.4 ± 0.2
5.8 ± 0.4
5.9 ± 0.0
CsrC-H91A Half-life
(min) 0 2 4 6 8 16 32
No C
Glc
Suc
Time (min)
CsrB CsrC
1.0 1.0±0.2 0.4±0.1 0.7±0 1.0 0.9±0.1 0.4±0.1 1.4±0.2
LB No C Glc Suc LB No C Glc Suc
Relative
level
59
Figure 2-10. EIIAGlc and MshH (CsrD ortholog) affect CsrB and CsrD decay in V.
cholerae. A, C and E) Decay rates of CsrB/C/D sRNAs were determined by Northern blotting, after rifampicin addition (time points indicated) to 27°C exponentially growing (OD600 ~0.5) cultures of V. cholerae 01 EI Tor (WT), Δcrr, ΔmshH and Δcrr ΔmshH strains. The RNA half-lives and standard derivations from duplicate experiments were determined as in Figure 2-1. B, D and F) Quantification of signals obtained from two independent experiments, as presented in panels A, C and E. Standard deviations are indicated.
A B
C D
E F
WT
Δcrr
ΔmshH
Δcrr ΔmshH
Time (min) 0 4 8 16 32
CsrB
14.0 ± 1.2
>32
>32
>32
Half-life (min)
CsrD
0 4 8 16 32
10.8 ± 2.0
>32
>32
>32
WT
Δcrr
ΔmshH
Δcrr ΔmshH
Time (min)Half-life
(min)
CsrC
>32
>32
>32
>32
WT
Δcrr
ΔmshH
Δcrr ΔmshH
Time (min) 0 4 8 16 32 Half-life
(min)
60
Figure 2-11. Proposed model for the effect of carbon availability on CsrB/C decay.
When glucose is present, EIIAGlc is mostly dephosphorylated, it binds to the EAL domain of CsrD and promotes CsrB/C decay. This results in increased CsrA activity to allow expression of genes and pathways needed for rapid growth, e.g. glycolysis. However, when preferred carbon resources have been expended and end-products have accumulated, the BarA-UvrY TCS activates CsrB/C synthesis and CsrD-dependent turnover is repressed. The resulting accumulation of CsrB/C will antagnize CsrA activity and promote the transition from glycolytic metabolism and active growth to gluconeogenesis, glycogen biosynthesis and the formation of a stress resistant phenotype.
UvrY
GGDEF
EAL
Glucose
Glu-6-P
HPr HPr-P
EI-P EI
PEP Pyruvate
P
P
CsrB/C
Carboxylic acids
mRNA translation, stability and transcription
elongation
CsrD
CsrA
Inner membrane
EIIAGlc EIIAGlc
EIIBGlc
EIICGlc
BarA RNase E
EIIAGlc
61
CHAPTER 3 EXPLORING THE MOLECULAR MECHANISM BY WHICH CsrD FACILITATES CsrB/C
TURNOVER
Introduction
In the last decade, the importance of sRNAs in gene expression has been
increasingly recognized in prokaryotes and eukaryotes (Gottesman & Storz, 2011, Storz
et al., 2011, Filipowicz et al., 2008, Vaucheret, 2006). While most of the bacterial
sRNAs act by base-pairing with mRNAs and altering gene expression, a small number
of sRNAs directly bind proteins and modulate protein activities by mimicking their
nucleic acid substrates. For example, Csr/Rsm sRNAs control gene expression and
cellular phenotypes indirectly by antagonizing CsrA/RsmA activities (Romeo, 1998,
Babitzke & Romeo, 2007, Romeo et al., 2013, Vakulskas et al., 2015). CsrA/RsmA are
RNA binding proteins that prefer to bind GGA motifs within apical loops of hairpin
structures in sRNAs or in the untranslated leader region or early coding region of target
mRNAs (Dubey et al., 2005, Holmqvist et al., 2016). CsrA/RsmA activate or repress
expression of target mRNAs by affecting the translation efficiency (Baker et al., 2002,
Baker et al., 2007, Dubey et al., 2003), transcription elongation (Figueroa-Bossi et al.,
2014), and/or transcript stability (Liu et al., 1995, Wang et al., 2005, Yakhnin et al.,
2013). In E. coli, CsrA activity is mainly controlled by the steady state levels of two non-
coding sRNAs CsrB and CsrC. These sRNAs contain many high-affinity CsrA binding
sites that sequesting CsrA from binding to its lower affinity target mRNAs (Liu et al.,
1997, Weilbacher et al., 2003).
Levels of CsrB/C are positively and negatively regulated by diverse regulators at
the synthesis level (Chavez et al., 2010, Huang et al., 2008, Edwards et al., 2011,
Suzuki et al., 2002, Camacho et al., 2015, Vakulskas et al., 2014, Zere et al., 2015,
62
Pannuri et al., 2016) and turnover level (Suzuki et al., 2006, Leng et al., 2016,
Vakulskas et al., 2016) in response to different environmental stimuli. This regulation
permits bacteria cell to fine-tune CsrA activity and gene expression for proper cellular
stress response.
Turnover of CsrB is mediated by RNase E and PNPase. The cleavage of CsrB is
initiated by RNase E at a 9 nt region located immediately upstream of intrinsic
terminator (Vakulskas et al., 2016). Deletion of this region or two adjacent adenine
residues within this region virtually eliminated CsrB turnover. PNPase acts to eliminate
the products from RNase E cleavage (Suzuki et al., 2006). In addition to RNase E and
PNPase, CsrD, EIIAGlc and CsrA are identified as key factors in regulating CsrB/C
turnover, as shown in Figure 4-1 (Suzuki et al., 2006, Leng et al., 2016, Vakulskas et
al., 2016). Deletion of CsrD significantly stabilized CsrB/C from half-lives of 1-2 min to
more than 32 min, indicating that CsrD is required for the rapid degradation of CsrB/C.
CsrD is not a nuclease (Suzuki et al., 2006), but is a membrane-bound signaling protein
containing GGDEF and EAL domains, which typically catalyze the synthesis and
turnover of the secondary messenger c-di-GMP. However, CsrD lacks critical catalytic
residues in these domains and displays no activity in c-di-GMP metabolism (Suzuki et
al., 2006). We recently showed that CsrD is responsible for the activation of CsrB/C
decay by EIIAGlc in the presence of a preferred carbon source (Fig. 3-1) (Leng et al.,
2016). When glucose is being actively transported by the PTS system,
unphosphorylated EIIAGlc accumulates and binds to the EAL domain of CsrD. Binding of
EIIAGlc allosterically activates CsrD activity to facilitate CsrB/C turnover (Leng et al.,
2016). Furthermore, CsrD and CsrA showed antagonistic roles on CsrB turnover (Fig. 3-
63
1). Specifically, the binding of CsrA to two GGA sites adjacent to the initial RNase E
cleavage region stabilized CsrB by protecting it from RNase E cleavage in a csrD
mutant and in vitro. However, CsrA disruption did not change CsrB decay rate in the
csrD WT strain (Vakulskas et al., 2016). In addition, even though CsrD is required for
the rapid degradation of CsrB in wild-type strain (Suzuki et al., 2006), it is not necessary
in csrA mutant strain, where CsrB is not covered by CsrA and can be rapidly degraded
by RNase E (Vakulskas et al., 2016). These data together suggest that CsrD has
evolved from a c-di-GMP metabolizing enzyme to become a device for decoupling Csr
sRNA turnover from the direct influence of CsrA binding.
So far, exactly how CsrD permits RNase E access to CsrB is not yet clear.
Previous study showed that the GGDEF domain of CsrD was homologous to several
RNA binding proteins, and CsrD bound to CsrB/C sRNAs in vitro, albeit nonspecifically
(Suzuki et al., 2006). Thus, we proposed that CsrD might directly bind CsrB/C, which in
turn releases CsrA and exposes the RNase E cleavage sites. Nevertheless, it is also
possible that CsrD works indirectly through other factors.
Materials and Methods
Media and Growth Conditions
Bacterial strains utilized in this study are listed in Table B-1. E. coli strains were
routinely grown in LB medium (1% tryptone, 1% NaCl, and 0.5% yeast extract) with
appropriate antibiotics when needed: ampicillin (100 μg mL−1), kanamycin (50 μg mL−1),
and gentamicin (10 g μg mL−1). For immunoprecipitation assays, overnight culture
grown in LB broth were inoculated into Kornberg medium (1.1% K2HPO4, 0.85%
KH2PO4, 0.6% yeast extract and 0.5% glucose) to an OD600 of 0.01, and their growth
was monitored at OD600. Strains carrying rne-1 were incubated at 30°C and then shifted
64
to 43-44°C before culture collection to inactive RNase E. Strains deficient in enolase
(PBAD-eno without arabinose) were grown in M9 medium containing 0.2% succinate and
0.02% glycerol.
Construction of Strains and Plasmids
E. coli gene deletions were created by the standard P1vir transduction procedure
or the lambda Red system as described (Datsenko & Wanner, 2000). The resistance
markers introduced into mutant strains were eliminated using an FLP expression
plasmid pCP20 when necessary (Datsenko & Wanner, 2000). The cat-PBAD-eno was
moved from TM447 (Morita et al., 2004) to MG1655 by P1 vir transduction.
Plasmids and DNA oligonucleotides used in this study are listed in Tables B-2
and B-3. For constructing plasmid p3FLAG-CsrD expressing CsrD containing a 3×FLAG
tag at C-terminal end, plasmid pBYH4 expressing WT-CsrD under the control of its
native promoter (Suzuki et al., 2006), was amplified with primer pair pBR3FLAG-F/R
(containing sequence encoding 3×FLAG tag), and the PCR product was re-ligated using
T4 DNA ligase (NEB). For plasmid p2SLStrep-CsrB expressing the Streptotag-CsrB,
plasmid pCsrB expressing the WT-CsrB under the control of its native promoter
(Vakulskas et al., 2016), was amplified with primer pair pBRStrep-CsrB-F/R (containing
Streptotag sequence), and PCR product was re-ligated.
Gel Mobility Shift Assay
RNAs (CsrB, CsrC, rpsT and GlmZ) were synthesized and end-labeled as
previously described (Suzuki et al., 2006). RNAs were gel purified, suspended in TE
buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA) and renatured by heating to 70°C for 5
min and slow cooling to room temperature. Binding reactions (10 μl) contained 10 mM
Tris-HCl, pH7.5, 125 mM KCl, and 2 mM MgCl2, 32.5 ng of yeast RNA, 7.5% glycerol, 1
65
mM dithiothreitol (DTT), 4 U of RNase inhibitor (Ambion), 0.6 nM CsrB RNA, purified
MBP-tagged CsrDΔTM, His-tagged EIIAGlc and/or His-tagged CsrA and 0.1 mg mL-1
xylene cyanol. For competition studies, assays were carried out with unlabeled RNA
competitors (CsrC, rpsT and GlmZ). Reaction mixtures were incubated for 30 min at
37°C to allow protein–RNA complex formation. Samples were then fractionated on 5%
native Bis-Tris gels. Radioactive bands were visualized and quantified using a
phosphorimager and Image Quant software.
In vitro RNase E Cleavage Assays
RNase E assays were performed by first incubating 0.05 nM RNA at 90°C for 3
min in RNase E reaction buffer (25 mM Tris–HCl, pH 7.9, 5 mM MgCl2, 60 mM KCl, 100
mM NH4Cl, 15 mM DTT and 7.5% glycerol), followed by cooling to 25°C over 10 min.
The reactions were then incubated at 25°C for 10 min in the presence or absence of
CsrA. RNase E was added to 37.5 nM and reactions were incubated at 25°C for an
additional 10 min. Reactions were stopped with two volumes of RNA loading buffer,
incubated at 65°C for 10 min to inactivate CsrA and subsequently kept on ice prior to
electrophoresis. Samples were separated by electrophoresis on 8% acrylamide gels
(19:1) containing 7 M urea for 3 h at 45 Watts. Gels were dried onto chromatography
paper and subjected to autoradiography.
Affinity Purification of CsrD and its Binding RNAs
Strains carrying p3FLAG-CsrD or pBYH4 were grown in 500 mL Kornberg
medium at 37°C, 250 rpm. At OD600 of 1.0, formaldehyde (0.5% final concentration) was
added and the cultures were incubated for 10 min at 30°C, 150 rpm. Glycine (0.125M
final concentration, pH 8.0) was subsequently added to quench the crosslinking reaction
and the samples were mixed with gentle swirling for 5 min at room temperature. The
66
cells were harvested by centrifugation (7K rpm, 10 min), washed twice with ice-cold 1x
PBS, suspended in 20 mL lysis buffer (20 mM HEPES pH 7.9, 150 mM KCl, 10%
glycerol, 0.5% Triton X-100) with EDTA-free protease inhibitor cocktail (Roche
Diagnostics) and lysed using 4 rounds of 10 sec sonication (power 30%, 0.5 sec pulse
on, 0.5 sec off) on ice. Lysates were cleared by centrifugation (14K rpm, 30 min) and
filtration over Millex-HV 0.45 m PVDF filter-units (Millipore) before mixing with 200 μL of
pre-washed ANTI-FLAG M2 beads (Sigma). Cell lysate with beads were rotated for 4
hours at 4°C to immunoprecipitate FLAG-tagged CsrD. The beads were then washed
three times with 4 mL high salt buffer (lysis buffer with 0.75M KCl), three times with 4mL
lysis buffer and once with elution buffer (20 mM HEPES pH 7.9, 150 mM KCl, 10%
glycerol). At last, CsrD-RNA complexes were eluted from the beads with 3X FLAG
Peptide (sigma), concentrated with Amicon Ultra-0.5 mL Centrifugal Filters (3kDa,
Millipore), and the formaldehyde crosslinking was reversed by heating at 95°C for 20
min. Afterward, RNAs were isolated by phenol chloroform extraction and ethanol
precipitation of the aqueous phase and proteins were isolated by acetone-precipitation
of the organic phase and dissolved in protein loading buffer. CsrB levels in samples
were analyzed by Northern blots and proteins were size-separated by 4–20% Mini-
PROTEAN ® TGX™ Precast gels (Biorad) and visualized by silver staining (kit from
Invitrogen).
Affinity Purification of in vivo Synthesized CsrB and its Associated Proteins
Strains carrying p2SLStrepto-CsrB or pCsrB were grown in 500 mL Kornberg
medium at 30°C, 250 rpm. At exponential phase of growth (OD600 of 0.5-0.6), cultures
were incubated at 50°C for ~15 min (temperature of culture reached 43-44°C) to
inactive RNase E. The cells were then harvested by centrifugation, washed once with
67
ice-cold 1x PBS, and suspended in 10 mL lysis buffer (10 mM Tris-HCl pH 7.8, 200 mM
KCl, 1 mM MgCl2, 5% glycerol, 0.5% Triton X-100) with EDTA-free protease inhibitor
cocktail (Roche Diagnostics) and SUPERase RNase Inhibitor (Thermo Fisher).
Afterward, Cells were lysed using 4 rounds of 10 sec sonication (power 30%, 0.5 sec
pulse on, 0.5 sec off) on ice. Lysates were then cleared by centrifugation (30 min, 14K
rpm) and filtration over Millex-HV 0.45 m PVDF filter-units (Millipore).
Dihydrostreptomycin (Sigma) coupled to epoxy-activated sepharose 6B (Sigma)
were prepared as previously described (Bachler et al., 1999, Windbichler & Schroeder,
2006) and used to pull down Strepto-CsrB. For preparation of the affinity column, 1 mL
slurry of the resin was applied to Poly-Prep® Chromatography Columns (BioRad). The
prepared column was washed twice with 6 mL lysis buffer. Subsequently, the cleared
bacterial lysate was loaded onto the column and incubated at 4°C for 3 hours on a
rotating wheel, followed by three washes with 6 mL washing buffer (10 mM Tris-HCl pH
7.8, 200 mM KCl, 1 mM MgCl2, 5% glycerol, 0.5% Triton X-100) and one time wash with
elution buffer (10 mM Tris-HCl pH 7.8, 200 mM KCl, 1 mM MgCl2, 5% glycerol). The
beads were eluted twice with 600 μL of freshly prepared 10 μM streptomycin in elution
buffer and elutes were concentrated with Amicon Ultra-0.5 mL Centrifugal Filters (3kDa,
Millipore) to 300 μL. RNA was extracted with phenol-chloroform and ethanol
precipitation of the aqueous phase. For protein isolation, the organic phase was
subjected to acetone precipitation and the pellet was dissolved in protein loading buffer.
Mass Spectrometry
For mass spectrometric analysis of CsrB associated proteins, precipitated
proteins were size-separated on a 4–20% Mini-PROTEAN ® TGX™ Precast gels
(Biorad) and observed with silver staining. Bands of interest were cut-out and the
68
excised gel pieces were destained with 50 mM ammonium bicarbonate, pH 8.5, and
then reduced by 10 mM Dithiothreitol (DTT, Sigma-Aldrich Inc., St. Louis, MO) at 37 ˚C
for 1h, followed by alkylation by 20 mM iodoacetamide in the dark for 30 min. Trypsin
(Sigma-Aldrich Inc.) was added for digestion (w/w for enzyme: sample = 1 : 100)
overnight at 37 °C. The digested peptides were desalted using micro ZipTip mini-
reverse phase (Millipore Inc., Billerica, MA), and then lyophilized to dryness.
Peptides derived from the protein samples and resuspended in 0.1% formic acid
for mass spectrometric analysis. The bottom-up proteomics data acquisition was
performed on an EASY-nLC 1200 ultra-performance liquid chromatography system
(Thermo Scientific Inc., Odense, Denmark) connected to an Orbitrap Fusion Tribrid
instrument equipped with a nano-electrospray source (Thermo Scientific Inc, San Jose,
CA). The peptide samples were loaded to a C18 trapping column (75 μm i.d. × 2 cm,
Acclaim PepMap® 100 particles with 3 μm size and 100 Å pores) and then eluted using
a C18 analytical column (75 μm i.d. x 15 cm, 3 μm particles with 100 Å pore size). The
flow rate was set at 300 nL/minute with solvent A (0.1% formic acid in water) and
solvent B (0.1% formic acid and 99.9% acetonitrile) as the mobile phases. Separation
was conducted using the following gradient: 2 - 35 % of B over 0 - 20 min; 35 - 98 % of
B over 20 – 21 min, and isocratic at 98% of B over 21-35 min. The full MS1 scan (m/z
350 - 2000) was performed on the Orbitrap with a resolution of 120,000 at m/z 400. Raw
data were analyzed using Mascot (Matrix Science, London, UK; version 2.4.1) against
E. coli K12 MG1655 20161109 database and Scaffold (version Scaffold_4.2.1,
Proteome Software Inc., Portland, OR) was used to validate MS/MS based peptide and
69
protein identifications. Protein identities were based on a threshold of 99.0 % probability
and < 0.1 % False Discovery Rate (FDR).
Protein Purification and Western Blots
Procedures for purification of His-tagged EIIAGlc and MBP-tagged CsrD and
Western blots were as described in Chapter 2 ‘Materials and Methods’. Purification of
RNase E and CsrA was as described previously (Vakulskas et al., 2016).
RNA Purification and Northern Blots
Total RNA from 20 μL resuspended intact cells, lysate and flow-through were
extracted using hot phenol chloroform (Georgellis et al., 1992) and purified with ethanol-
precipitation. Procedures for Northern blots were as described in Chapter 2 ‘Materials
and Methods’.
Results and Discussion
EIIAGlc is not Capable to Stimulate the Binding of CsrD to CsrB in vitro
A recent study indicated that EIIAGlc directly binds CsrD and allosterically
stimulates its activity (Leng et al., 2016). We therefore hypothesized that EIIAGlc might
promote the binding between CsrD and CsrB if CsrD functions as a CsrB binding
protein. To verify this, we compared the binding affinities and specificities of CsrD to
CsrB in the presence and absence of EIIAGlc using EMSA. Fig. 3-2A showed that CsrD
binds CsrB in vitro, but the binding affinity between CsrD and CsrB is much lower than
that between CsrA and CsrB (Kd value of 0.8-1.6 nM) (Weilbacher et al., 2003). Addition
of EIIAGlc did not alter the binding affinity (Fig. 3-2B). The specificity of the CsrD-CsrB
interaction was investigated by performing competition experiments with specific
(sRNAs CsrB and CsrC) and non-specific (mRNA rpsT and sRNA GlmZ) unlabelled
RNA competitors (Fig. 3-2C). We found that while CsrB, CsrC and rpsT transcripts
70
competed for CsrD binding, non-specific sRNA GlmZ did not compete (Fig. 3-2C),
suggesting that CsrD binds to CsrB with some specificitiy. Moreover, the specificity of
binding was not altered by addition of EIIAGlc (Fig. 3-2D). This suggests that CsrD binds
to CsrB with some specificitiy and EIIAGlc did not alter the specificity of this binding. All
together, EIIAGlc does not promote the binding affinity or specificity of CsrD to CsrB in
vitro.
CsrA Influences the CsrD-CsrB Interaction in vitro
Another recent study revealed that CsrD is required for CsrB turnover when the
RNase E-mediated CsrB cleavage can be protected by CsrA (Vakulskas et al., 2016).
Then, it is possible that CsrD prefers to bind CsrB in complex with CsrA. Specifically,
binding of CsrA to CsrB may modulate the sencondary or tertiary structure of CsrB and
render it accessible to CsrD binding. To test this, we examined the effect of CsrA on the
binding of CsrD to CsrB using EMSA. As expected, the binding of CsrA to CsrB was
observed in the absence of CsrD (the last lane, Fig. 3-3B). As the concentration of CsrD
increased, an additional shift was observed (Fig. 3-3B). Nevertheless, this shift does not
correspond to the CsrD-CsrB complex shown in Fig. 3-3A, where CsrA is not present.
Addition of EIIAGlc to the binding reaction did not alter the shift pattern (Fig. 3-3C). This
data suggest that addition of CsrA influences the CsrD-CsrB binding complex. We
speculate that the additional shift in Fig. 3-3B might represent a complex formed by
CsrD, CsrB and CsrA, but this needs to be further investigated by western blot analysis.
CsrD cannot Facilitate CsrA-mediated Protection of CsrB Cleaved by RNase E
CsrB contains 22 GGA motifs, most of which are presumed to be CsrA binding
sites (Liu et al., 1997, Weilbacher et al., 2003). Although the binding affinity of CsrD with
CsrB is much lower than that of CsrA with CsrB (Fig. 3-2A) (Weilbacher et al., 2003), we
71
cannot rule out the possibility that CsrD competes with CsrA for binding to the two GGA
motifs adjacent to the RNase E cleavage site, and expose CsrB for RNase E cleavage.
Thus, we examined whether CsrD facilitates the in vitro cleavage of CsrA-protected
CsrB by RNase E. This assay was performed with a shorter fragment of CsrB (+226 to
+369 relative to the transcription start site), whose turnover remains CsrD-dependent.
Our data indicated that RNase E cleaves the CsrB fragment mainly at nucleotides A327
and A331, which are located within the RNase E initial cleavage region ‘A’ (Vakulskas et
al., 2016). Addition of CsrA completely blocked the cleavage of CsrB by RNase E at two
major cleavage sites. However, inceasing concentration of CsrD with the help of EIIAGlc
did not facilite the cleavage of CsrA-protected CsrB by RNase E (Fig. 3-4). This
suggests that rather than functioning via a direct binding to CsrB, CsrD might require
other factor(s) in addition to EIIAGlc to render the CsrA-protected CsrB accessible to
RNase E cleavage.
CsrD is Unlikely to Bind CsrB in vivo
We further tested the in vivo binding of CsrD to CsrB using formaldehyde
crosslinking and immunoprecipitation, which could potentially capture transient and
unstable binding. If CsrD binds to CsrB in vivo, CsrB is expected to be enriched along
with FLAG-tagged CsrD. A 3×FLAG tag was added to the C terminus of CsrD, which
does not affect the activity of CsrD on CsrB turnover (Fig. 3-5A). FLAG-tagged CsrD
was expressed under the control of its native promoter on a multi-copy plasmid pBR322
in a strain with a csrD genomic deletion. Plasmid pBYH4 expressing untagged CsrD
was used as the negative control. The crosslinked CsrD-RNA complex was captured
and subsequently uncrosslinked by heating at 95 °C for 20 min. Enrichment of CsrD and
CsrB was determined by silver staining and Northern blot, respectively. Fig. 3-5B
72
showed that FLAG-tagged CsrD was enriched in samples expressing tagged CsrD
comparing to the negative control (lane 1 and 2). Samples without formaldehyde
treatment (lane 3) gave a stronger enrichment of FLAG-tagged CsrD, suggesting that
formaldehyde treatment somehow inhibited the binding of FLAG-tagged CsrD to the
beads. Surprisingly, while CsrD was strongly enriched, no enrichment of CsrB was
detected (Fig. 3-5C). These data suggest that CsrD might not directly bind to CsrB in
vivo. Although it is also possible that binding between CsrD and CsrB is very transient
and could not be captured with the approach we used.
CsrB Associated Proteins and their Influence on CsrB Turnover
To further investigate binding of CsrD with CsrB in vivo, we used a reciprocal
approach, by pulling down proteins from the cell via a tagged CsrB. This approach will
also provide information on the proteins associated with CsrB and may help to identify
the missing factor(s) involved in CsrB/C turnover.
Our previous work showed that the second stem loop of CsrB was not essential
for the CsrD-mediated turnover of CsrB (Vakulskas et al., 2016). This stem loop was
partially substituted with an RNA aptamer Streptotag that specifically binds to
streptomycin with high binding affinity (Bachler et al., 1999, Dangerfield et al., 2006).
Tagged CsrB was expressed under the control of its own promoter on plasmid pBR322
in a strain lacking the endogenous csrB. CsrB decay assays revealed that the RNA
aptamer did not influence the expression of CsrB, and slightly stabilized CsrB from a
half-life of 4.3 min to 8.3 min (Fig. 3-6A). Nevertheless, decay of the tagged CsrB
remains CsrD dependent (Fig. 3-6A). We tried to capture the transient CsrB-protein
complex with formaldehyde crosslinking, but this treatment severely inhibited the
binding of tagged CsrB to the beads (data not shown). Then we performed this
73
experiment using strains in rne-1 background and the bacterial cultures were shifted
from 30°C to 43-44°C prior to affinity purification to inactive RNase E. Under this
condition, CsrB-protein complex could potentially be stabilized. After affinity purification,
enrichment of CsrB and CsrD was analyzed by Northern blot and Western blot,
respectively. Northern blot results illustrated that while 10% of the total CsrB was
captured using this approach (data not shown), CsrD was not recovered by CsrB,
further supporting our previous finding that CsrD might not directly bind CsrB in vivo
(Fig. 3-6C). CsrB associated proteins were separated on SDS-PAGE gel and observed
with silver staining, as shown in Fig. 3-6B. Of particular interest, in addition to CsrA, the
well-characterized CsrB binding protein, two other bands A and B showed strong
enrichments in the strain expressing tagged CsrB (lane 1 and 2) relative to the negative
control with the wild-type CsrB (lane 3). Using LC-MS/MS, we analyzed all the proteins
from the two bands, as shown in Tables 3-1 and 3-2. Unexpectedly, no protein from
band A was identified to have a size corresponding to band A. In band B, enolase
represents the most abundant protein and has a size matching that of band B. Enolase
is a glycolytic enzyme that catalyzes the interconversion of 2-phospho-D-glycerate and
phosphoenolpyruvate and is a component of the RNA degradosome. The role of
enolase in RNA degradosome is not very clear, but it is required for the destabilization
of ptsG mRNA in response to glucose-6-phosphate (Morita et al., 2005, Morita et al.,
2004). Given that enolase in eukaryotes is capable to bind RNA (Hernández-Pérez et
al., 2011), we hypothesized E. coli enolase directly binds CsrB and facilitates its
turnover. To examine the possible role of enolase in CsrB turnover, we compared CsrB
decay rates in wild-type strain and a mutant strain growing in M9 minimal medium
74
supplemented with succinate and glycerol, in which the eno gene is under the control of
an arabinose-inducible promoter PBAD (Morita et al., 2004). In the absence of arabinose,
enolase should not be expressed and the decay rate of CsrB was moderately
decreased in enolase deficient strain (half-life of 8.2 min) as compared to in the wild-
type strain (half-life of 5.0 min) (Fig. 3-7A). Since the bacterial culture used for
immunoprecipitation assay was grown in Kornberg medium (Fig. 3-6), we also
examined the effect of enolase on CsrB turnover in Kornberg medium. We first
transferred the exponentially growing culture in M9 minimal medium to Kornberg
medium and then compared CsrB decay rates in enolase deficient strain and wild-type
strain 10 min after the transfer. CsrB decay rates were faster in Kornberg medium than
in the minimal medium with succinate and glycerol likely due to the presence of glucose
(Fig. 3-7A and B). However, decay rates of CsrB were essentially identical in strains
lacking enolase (half-life of 2.3 min) or with enolase (half-life of 2.2 min), suggesting that
enolase has little or no role in CsrB turnover (Fig. 3-7B). Whether enolase directly binds
to CsrB remains to be further determined, but based on our finidng that enolase has no
effect on CsrB turnover and it directly interacts with RNase E (Mackie, 2013), it is
possible that enolase associates with CsrB indirectly via RNase E.
Meanwhile, we also observed other dramatically enriched proteins in amounts
less abundant than enolase and with sizes that do not correspond to bands A and B
(Tables 3-1 and 3-2). They are represented by RNase E, poly(A) polymerase (PAP I),
RhlB and multiple ribosomal proteins. It is known that RNase E directly binds CsrB and
cleaves it. RhlB, the component of of RNA degradosome comprising of RhlB, PNPase,
enolase and RNase E, could have been recruited by RNase E. PAP I promotes
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degradation of sRNA and mRNA by adding a single stranded poly(A) tail at the 3’ end of
the RNAs (Xu et al., 1993, Hajnsdorf et al., 1995), but it does not influence CsrB
turnover (Suzuki et al., 2006). It is also reported to associate with PNPase and might
have been recuirted via the RNA degradosome (Mohanty et al., 2004). Ribosomal
proteins were capable to interact directly with RNase E (Tsai et al., 2012), so they might
directly bind the multiple AGGA motifs within CsrB, or indirectly associate with CsrB via
its interaction with RNase E. Overall, copurification of these proteins suggest that CsrB
might provide a platform for binding of a specific set of proteins, but whether these
proteins are functionally relevant to CsrB decay or have other functions remains to be
further explored.
Conclusion
Previous study demonstrated that CsrD facilitates CsrB turnover by overcoming
the CsrA mediated protection and promoting the initinal cleavage of CsrB by RNase E.
This study suggested that CsrD might not work as a RNA binding protein to render
CsrA-protected CsrB accessible to RNase E cleavage. In vitro assays suggest that
other factor(s) in addition to EIIAGlc and CsrA might be required to transmit the
regulatory role of CsrD to CsrB turnover. In addition, proteins associated with CsrB were
detected in this study, but whether these interactions have any physiological functions,
such as affecting CsrB turnover, needs to be further determined.
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Figure 3-1. CsrB decay is regulated in response to carbon availability through its effect
on CsrA and CsrD antagonism (Vakulskas et al., 2016). The phosphorylation state of the PTS protein EIIAGlc serves as an indicator of carbon availability. P-EIIAGlc predominates when a preferred carbon source such as glucose is unavailable, and this form is unable to bind to CsrD. During glucose transport, EIIAGlc becomes dephosphorylated and able to bind to CsrD and potentiate CsrB decay. CsrA binding to CsrB protects it from RNase E cleavage in the absence of CsrD-EIIAGlc. A broken line indicates that the molecular mechanism of CsrD remains to be determined. RNase E, PNPase and other nucleases degrade CsrB to nucleotides (NTDs).
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Figure 3-2. EIIAGlc has no effect on the binding of CsrD to CsrB RNA in vitro. A and B)
EMSA experiments were carried out with 0.6 nM labeled CsrB and increasing concentrations of CsrD (0-320 nM) in the absence or presence of EIIAGlc. C and D) Competition assays were performed using 0.6 nM labeled CsrB with indicated concentration of CsrD and EIIAGlc in the absence or presence of 100-fold and 1,000-fold molar excess of unlabeled competitor sRNAs, CsrB, CsrC, and GlmZ and mRNA, rpsT.
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Figure 3-3. CsrA influences the CsrD-CsrB complex in vitro. EMSA experiments were
carried out with 0.6 nM labeled CsrB and increasing concentrations of CsrD (0-320 nM) in the absence or presence of CsrA and EIIAGlc.
0 10 20 40 80 120 160 250 320 CsrD (nM) 0 10 20 40 80 120 160 250 320 0 CsrD (nM)
CsrA (nM) 0 500
A B
C
0 10 20 40 80 120 160 250 320 0 CsrD (nM)
CsrA (nM) 0 500
EIIAGlc(nM) 0 1280
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Figure 3-4. Effect of EIIAGlc and CsrD on RNase E-dependent cleavage of CsrB in vitro.
This experiment was performed with the +226 CsrB RNA (CsrB alleles starting from nucleotide +226 of CsrB to the end of csrB gene), 0.5 mM CsrA, 37.5 nM RNase E, EIIAGlc (0.5, 2 or 6 μM), and CsrD (0.125, 0.5 or 1.5 μM) as indicated. Premixed EIIAGlc-CsrD complex or each protein individually was added to reactions and incubated at 25 °C for 10 min. CsrA and RNase E were then sequentially added to reactions and cleavage was allowed to proceed for 10 min at 25 °C. Control experiments were performed with RNA only. Partial alkaline hydrolysis (OH) and RNase T1 digestion (T1) ladders are also indicated. Numbering is with respect to the full length CsrB sequence.
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Figure 3-5. CsrB is not copurified with FLAG-tagged CsrD. A) Northern blot depicting
the CsrB decay rates in MG1655 ΔcsrD strains carrying pBYH4 or p3FLAG-CsrD, which express WT CsrD and FLAG-tagged CsrD, respectively. B) Proteins co-purifying with FLAG-tagged CsrD were observed by silver staining. 1, MG1655 ΔcsrD carrying pBYH4 with formaldehyde crosslinking; 2, MG1655 ΔcsrD carrying p3FLAG-CsrD with formaldehyde crosslinking; 3, MG1655 ΔcsrD carrying p3FLAG-CsrD without formaldehyde crosslinking. C) Northern blot depicting CsrB levels before, during and after the affinity purification. RNA samples were prepared from intact cells before lysate preparation (Tot), from the cleared lysate (Lys), from the flow-through fraction (FT) and from the eluates (Elu1 and Elu2). The amount of RNA loaded in Tot, Lys, FT each corresponds to 30 μL of bacterial culture, in Elu1 and Elu2 correspond to 4.5 mL and 60 mL of bacterial culture, respectively.
1 2 3 1 2 3 1 2 3 1 2 3 1 2 3
Tot Lys FT Elu1 Elu2
A
B
C
CsrDFLAG
1 2 3
0 2 4 8 16 0 2 4 8 16 Time (min)
CsrDWT CsrDFLAG
CsrB
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Figure 3-6. CsrD is not recovered by Strepto-CsrB in vivo. A) Streptotag does not affect
expression of CsrB and the CsrD-mediated turnover of CsrB. Rifampcin was added to exponentially growing cultures of MG1655 ΔcsrB and MG1655 ΔcsrB ΔcsrD and decay rates of CsrB were determined by Northern blots as described in Chapter 2 ‘Materials and Methods’. B) Proteins co-purifying with Strepto-CsrB were observed by silver staining. 1, MG1655 rne-1 ΔcsrB harboring p2SLStrep-CsrB; 2, MG1655 rne-1 ΔcsrB ΔcsrD harboring p2SLStrep-CsrB; 3, MG1655 rne-1 ΔcsrB harboring pCsrB. C) Western blot depicting CsrD levels before, during and after the affinity purification. Proteins samples were prepared from intact cells before lysate preparation (Tot), from the cleared lysate (Lys), from the flow-through fraction (FT) and from the eluates (Elu1 and Elu2). The amount of proteins loaded in Tot, Lys, FT each corresponds to 250 μL of bacterial culture, in Elu1 and Elu2 correspond to 16.5 mL and 250 mL of bacterial culture, respectively.
CsrB
1 2 3
A?
B?
CsrA
kDa
75
50
37
25 20 15
100
A
C
B
0 2 4 8 16 0 2 4 8 16
0 4 8 16 32 0 4 8 16 32
WT-CsrB Strepto-CsrB
WT-CsrB Strepto-CsrB
csrB
csrB csrD
Half-lives
Half-lives
4.3 min 8.3min
>32min >32min
ΔcsrD WT-CsrB Strepto-CsrB
Tot Tot Lys FT Elu1 Elu2 Tot Lys FT Elu1 Elu2
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Table 3-1. Proteins co-purifying with Strepto-CsrB identified from band A by mass-spectrometry
Protein Gene MW (kDa)
Ratio of Total Unique Spec
Counts
Coverage (%)
Enolase eno 46 3 6.5 RNase E rne 118 3 13
RNA helicase RhlB
rhlB 47 3 7.4
50S ribosomal protein L2
rplB 30 3 17
50S ribosomal protein L3
rplC 22 5 23
50S ribosomal protein L15
rplO 15 6 40
50S ribosomal protein L15
rplI 16 4 35
50S ribosomal protein L21
rplU 12 4 37
50S ribosomal protein L1
rplA 25 3 17
30S ribosomal protein S3
rpsC 26 3 18
30S ribosomal protein S5
rpsE 18 4 34
Protein identities were based on a thresholdof 99.0 % probability and < 0.1 % False Discovery Rate (FDR).
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Table 3-2. Proteins co-purifying with Strepto-CsrB identified from band B by mass-spectrometry
Protein Gene MW (kDa)
Ratio of Total Unique Spec Counts
Coverage (%)
R1 R2 R1 R2
Enolase eno 46 3 9 41 64 Poly(A) polymerase
I pcnB 54 5 16 12 27
RNase E rne 118 8 7 8 8 50S ribosomal
subunit protein L9 rplI 16 3 10 40 61
50S ribosomal subunit protein L3
rplC 22 4 3 17 30
50S ribosomal subunit protein L21
rplU 12 3 5 25 32
Experiment was done in replicate. R1, repeat 1; R2, repeat 2. Protein identities were based on a thresholdof 99.0 % probability and < 0.1 % False Discovery Rate (FDR)
Figure 3-7. Enolase has little or no effect on CsrB turnover. Decay rates of CsrB were
determined by Northen blotting of RNA from E. coli MG1655 (WT) and PBAD- eno strains. A) Rifampicin was added to bacterial cultures growing in M9 minimal medium containing 0.2% succinate and 0.02% glycerol entering exponential phase. B) Strains were grown in M9 minimal medium containing 0.2% succinate and 0.02% glycerol to exponential phase, washed and reinoculated into Kornberg medium. Rifampicin was added 10 min after the reinoculation. RNA half-lives were determined as in Figure 2-1.
WT
0 2 4 6 8 16 32 Time (min) Half-lives (min)
5.0
8.2
WT
0 2 4 6 8 16 32 Time (min) Half-lives (min)
2.2
2.3
A
B
PBAD-eno
PBAD-eno
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CHAPTER 4 EPISTASIS ANALYSIS USING RNA-SEQ (EPI-SEQ) TO EXPLORE THE REGULATORY ROLE OF CsrD
Introduction
The Csr (Rsm) system is a global regulatory system that is conserved among
Gammaproteobacteria (Vakulskas et al., 2015, Zere et al., 2015). It controls complex
phenotypes, including glycogen metabolism (Baker et al., 2002, Romeo et al., 1993),
biofilm formation (Jonas et al., 2008, Sterzenbach et al., 2013, Wang et al., 2005),
motility (Wei et al., 2001, Yakhnin et al., 2013), and virulence (Bhatt et al., 2009,
Martínez et al., 2011, Vakulskas et al., 2015). The Csr system in E. coli consists of the
RNA binding protein CsrA, the inhibitory small RNAs (sRNAs) CsrB and CsrC, and a
specificity factor for the turnover of the sRNAs CsrD (Fig. 4-1A) (Vakulskas et al., 2015).
Generally, CsrA binds to conserved GGA motifs in the 5’-untranslated or early coding
region of mRNAs leading to changes in RNA structure (Patterson-Fortin et al., 2013),
RNA stability (Liu et al., 1995, Wang et al., 2005, Yakhnin et al., 2013), translation
initiation (Baker et al., 2002, Baker et al., 2007, Dubey et al., 2003), and/or transcription
elongation (Figueroa-Bossi et al., 2014). CsrA activity is controlled primarily by the
steady state levels of CsrB/C, which contain many high affinity CsrA binding sites that
sequester CsrA from interacting with its lower affinity mRNA regulatory targets (Liu et
al., 1997, Weilbacher et al., 2003, Vakulskas et al., 2015). Transcription of these sRNAs
is controlled by the BarA-UvrY TCS in response to the accumulation of end products of
metabolism, including acetate and formate (Chavez et al., 2010, Zere et al., 2015).
RNase E-dependent turnover of CsrB/C is regulated by CsrD although the exact
molecular mechanism remains unclear (Fig. 4-1A) (Suzuki et al., 2006, Vakulskas et al.,
2016, Leng et al., 2016). CsrD is a membrane-bound protein that has degenerate
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GGDEF and EAL domains. Although these domains are often involved in the synthesis
and degradation of c-di-GMP, CsrD neither produces nor degrades c-di-GMP (Suzuki et
al., 2006). However, CsrD is essential for the turnover of CsrB/C (Suzuki et al., 2006,
Vakulskas et al., 2016). Loss of CsrD strongly stabilizes these RNAs but only leads to a
modest increase in their steady states levels, likely due to regulatory feedback loops
that also control their transcription (Fig. 4-1A) (Suzuki et al., 2006).
When CsrD was initially described as a regulator of CsrB/C turnover, it was also
found to regulate CsrA-dependent gene expression and phenotypes such as biofilm
formation and glycogen synthesis in a CsrA and CsrB/C dependent manner (Suzuki et
al., 2006). This led to the development of a model, in which CsrD affects gene
expression through changes in CsrB/C levels, which then affect CsrA activity (Model 1,
Fig. 4-1A). According to this model CsrA acts as the most downstream regulator of gene
expression in the Csr system. It directly regulates gene expression posttranscriptionally
by affecting transcription elongation and transcript stability, but it can also mediate
indirect transcriptional and posttranscriptional effects through regulators it controls
directly. However, a recent transcriptomics study found that in addition to global effects
of CsrA on RNA stability and steady state RNA levels, CsrD had many effects on RNA
levels but not stability (Esquerré et al., 2016). These results led the authors to propose
an alternative model (Model 2, Fig. 4-1B). This model proposes that CsrA acts as a
posttranscriptional regulator for its direct targets, but it mediates most of its indirect
effects on RNA abundance indirectly though transcriptional effects controlled by CsrD
and perhaps some additional effects through other unknown factors.
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In this work we used RNA-seq to untangle the epistatic relationships among the
members of the Csr system (Epi-seq). To determine on a genome-wide scale whether
CsrD affects gene expression primarily through its effects on CsrB/C levels and
therefore CsrA activity, we compared the impact of CsrD on transcriptomic profile in the
presence and absence of CsrA or CsrB/C. If most of the effects of CsrD on gene
expression were lost in the absence of CsrA or CsrB/C, this would support Model 1 in
Fig. 4-1A, where CsrD primarily affects gene expression through CsrB/C and CsrA.
Likewise, CsrA effects on RNA abundance were assessed in strains with and without
CsrD. If most of CsrA effects on transcript levels were eliminated in the absence of
CsrD, this would support Model 2 in Fig. 4-1B, where CsrA primarily acts through CsrD
to mediate changes in transcription. We found that CsrA and CsrB mediate vast
transcriptional changes independently of CsrD, and the majority of CsrD-dependent
effects on gene expression require CsrA and CsrB/C. These results support Model 1,
where CsrD works primarily upstream of and through CsrB/C to regulate CsrA activity,
which is the major regulator of gene expression in the Csr system.
Materials and Methods
Media and Growth Conditions
Bacterial strains utilized in this study are listed in Table B-1. E. coli strains were
routinely grown in LB medium (1% tryptone, 1% NaCl, and 0.5% yeast extract) with
appropriate antibiotics when needed: ampicillin (100 μg mL−1), kanamycin (50 μg mL−1),
chloramphenicol (25 μg mL−1), and gentamicin (10 μg mL−1). For growth curve and
RNA-seq analysis, overnight culture grown in LB broth were inoculated into Kornberg
medium (1.1% K2HPO4, 0.85% KH2PO4, 0.6% yeast extract and 0.5% glucose) to an
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OD600 of 0.01, cultures were then grown at 37°C with shaking at 250 rpm and their
growth was monitored at OD600.
Construction of Strains and Plasmids
MG1655 pgaC880::cam, in which pagC gene was disrupted by transposon mini-
Tn10cam, was used as a parent wild-type strain (Wang et al., 2004). E. coli gene
deletions and disruptions were created by vir transduction procedure using E. coli donor
strains from previous studies (Suzuki et al., 2006, Wang et al., 2004, Vakulskas et al.,
2016) and the Keio library (Baba et al., 2006), as shown in Table B-1. The FRT-flanked
antibiotic resistance cassettes introduced into mutant strains were eliminated using an
FLP expression plasmid pCP20 when necessary (Datsenko & Wanner, 2000).
Plasmids and DNA oligonucleotides used in this study are listed in Tables B-2
and B-3. Plasmids p2VR112 (referred as pCsrA in this study) and pBRY4 (referred as
pCsrD in this study), express genes csrA and csrD, respectively, under the control of
their own promoters on plasmid pBR322 (Suzuki et al., 2006, Vakulskas et al., 2016).
To construct plasmid pCsrB for expression of CsrB, the csrB gene with 494 base pairs
(bp) upstream and 36 bp downstream was amplified from the genomic DNA and cloned
into plasmid pBR322. Control strains for pCsrA, pCsrD or pCsrB containing strains were
transformed with pBR322.
Glycogen Biosynthesis
Glycogen production was examined by staining colonies with iodine vapor (Liu et
al., 1997)
RNA Extraction and Purification
During transition to stationary phase of growth (OD600 of 2.0), 1 mL of cell culture
was collected and immediately mixed with 0.125 mL of stop solution (10% phenol / 90%
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ethanol) to stabilize RNA. Total RNA was extracted using hot phenol chloroform
followed by ethanol precipitation (Georgellis et al., 1992). Genomic DNA was removed
by treating 20 μg of nucleic acid with 4U of Turbo DNase (Ambion), and RNA was
purified from these reactions with the RNeasy kit (Qiagen). The integrity of the RNA was
verified using denaturing gel electrophoresis and RNA Bioanalyzer (Agilent) analysis.
Northern Blotting
Procedures for Northern blots were as described as in Chapter 2 ‘Materials and
Methods’.
RNA-seq Library Preparation
For each strain, independently grown biological triplicates were collected.
Ribosomal RNA was depleted from 5 μg of total RNA using the Ribo-Zero rRNA
Removal Kit for Gram-Negative Bacteria (Illumina). The concentrations of the rRNA-
depleted samples were determined with the Qubit RNA HS Assay Kit (ThermoFisher).
RNA-seq libraries were then generated with the Stranded RNA-Seq Library Preparation
Kit for Illumina (KAPA) and NEBNext Multiplex Oligos for Illumina adaptors (NEB)
according to the manufacturer’s instructions for 100 ng of starting material and a mean
insert size of 200-300 bases. Final libraries were purified with Pure Beads (KAPA).
Sequencing library size and integrity were verified with DNA Bioanalyzer analysis
(Agilent). Libraries were pooled and sequenced on 2 lanes of 50SE HiSeq 2500
(Illumina) by the Genomic Services Laboratory at the HudsonAlpha Institute for
Biotechnology.
RNA-seq Data Analysis
Raw reads were demultiplexed and analyzed with FastQC to ensure there were
no quality control issues. Sequencing reads were mapped to the E. coli rRNA
89
sequences with Bowtie 2 (Langmead & Salzberg, 2012), and unmapped reads were
retained. rRNA depleted reads were then mapped to the E. coli genomic DNA sequence
(NC_000913.3) with Bowtie 2 (Langmead & Salzberg, 2012). Read counts per gene
were calculated with htseq-count (Anders et al., 2015). Read counts were filtered to
remove genes with an average of 10 reads per sample across all samples. Differential
expression was analyzed with limma voom (Liu et al., 2015), and genes with fold
changes greater than 2 and a p-value less 0.05 were considered significant.
qRT-PCR
Quantitative reverse transcriptase PCR (qRT-PCR) was conducted using iTaq
Universal SYBR Green One-Step Kit (Bio-Rad) and an iQ5 iCycler real time PCR
system (Bio-Rad) according to the manufacturer’s instructions. Reactions of 10 μL
contained 200 ng of RNA or DNA standard, 300 nM of each primer, iScript reverse
transcriptase, and 1x iTaq universal SYBR Green reaction mix. Reactions were
incubated for 10 min of RT at 50°C, 1 min of denaturation and RT inactivation at 95°C,
and then 45 cycles of 10 sec of denaturation at 95°C and 20 sec of annealing,
extension, and imaging at 60°C. Melt curve analysis was used to verify the specificity of
amplifications with the parameters: 95°C for 1 min, 55°C for 1 min, and increasing the
temperature 0.5°C/10 sec until reaching 95°C. RNA abundances were determined
relative to a standard curve of PCR products and normalized to 16s rRNA levels.
Results and Discussion
Construction and Characterization of Bacterial Strains
The csrA::gm gene disruption mutant (hereafter csrA mutant) used in this study is
not a deletion, but expresses a functionally impaired CsrA protein (Vakulskas et al.,
2016). Unlike a csrA deletion strain which exhibits a severe growth defects and rapidly
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accumulates suppressor mutations (Lawhon et al., 2003, Timmermans & Van Melderen,
2009, Yakhnin et al., 2013), this csrA mutant grows similarly to the wild-type in rich
media. For example, during the transition to stationary phase of growth (OD600 of 2.0) in
Kornberg (KB) media where we collected RNA for this study, the csrA mutant grows
only slightly slower than the csrA wild-type strain (Fig. 4-2A). As a csrAD double mutant
showed dramatically enhanced cell aggregation and biofilm formation (data not shown),
all of the strains were constructed in a pgaC disruption mutant background, which
cannot synthesize PGA or form PGA-dependent biofilm (Wang et al., 2004).
Northern blots and quantitative reverse transcriptase PCR (qRT-PCR) (Figs. 4-
2B-D) showed that while CsrD is essential for normal turnover of CsrB/C (Suzuki et al.,
2006), deletion of csrD only moderately increased CsrB levels and slightly decreased
CsrC levels and this phenotype can be complemented by ectopic expression of csrD
(Figs. 4-2B-D). Likely this is due to a feedback loop within Csr regulatory circuitry, in
which CsrB antagonizes CsrA activity to repress CsrB and CsrC transcription through
the BarA-UvrY TCS (Fig. 4-1A). CsrB is the principal sRNA antagonist of CsrA. Thus,
we expect that even with a minor decrease in CsrC level, the increased CsrB level
caused by the csrD deletion is sufficient to reduce CsrA activity. As expected (Camacho
et al., 2015), disruption of csrA in either the csrD mutant or wild-type strains significantly
reduced CsrB/C levels via BarA-UvrY TCS and this reduction can be restored by
ectopic expression of csrA (Figs. 4-2 B-D). Overall, these observations are consistent
with the known regulatory circuitry outlined in Model 1, where multiple feedback loops
existing to regulate the levels of CsrB/C.
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We also used iodine staining and qRT-PCR to determine glycogen abundance
and glgC RNA transcript levels in all the strains. As expected, the csrA mutant
accumulated much more glycogen and significantly increased glgC expression 8.7-fold
relative to the wild-type strain (Figs. 4-2E and F), whereas deletion of csrD resulted in a
slight increase in glycogen levels and increased glgC expression 1.4-fold relative to the
wild-type strain (Fig. 4-2E and F). Ectopic expression of csrA in the csrAD mutant and
csrBC in csrBCD significantly reduced and stimulated glycogen synthesis, respectively.
The analysis of glgC mRNA levels as determined by qRT-PCR were also consistent
with these observations of glycogen levels (Fig. 4-2F). All of these results confirmed that
the strains we constructed for RNA-seq analysis behave as expected.
CsrA Retains its Global Role in Regulating Transcript Levels in the Absence of CsrD
Differential expression analysis revealed 1,054 genes with greater than 2-fold
change in RNA levels and p-values less than 0.05 between the wild-type and csrA
mutant strains (Fig. 4-3A). CsrA repressed the expression of 828 genes and activated
226. In addition, 807 out of the 1054 genes were differentially expressed between the
csrAD double mutant strain and the csrD mutant strain (csrAD-csrD, Fig. 4-3B),
suggesting that CsrA retains its global influence on 80% of its target genes in the
absence of CsrD. To examine whether CsrD influences CsrA-mediated changes in gene
expression, we compared the log2 transformed fold change in RNA abundance caused
by the csrA mutation in the wild-type (csrA - WT) and csrD mutant strain (csrAD - csrD)
backgrounds. The high value of Spearman’s correlation coeficient between these data
sets demonstrated that the absence of csrD had little impact on the overall effect of
CsrA on gene expression (ρ=0.78, Fig. 4-3C). In addition, few genes varied in
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expression in the presence or absence of CsrD (Fig. 4-3C). Similarly, ectopic
expression of csrA in the csrAD double mutant resulted in vast changes in gene
expression (csrAD pCsrA - csrAD, Fig. 4-3B). The log2 transformed fold changes from
this comparison showed a strong negative correlation with those resulting from mutation
of csrA in either a wild-type (csrA - WT) or csrD mutant strain (csrAD - csrD)
background (ρ=0.82 and 0.87, respectively, Figs. 4-3D and E). Together these data
indicate that CsrA does not require CsrD to exert global effects on transcript levels. The
data do not support Model 2 in which CsrA effects on the transcriptome are primarily
mediated through CsrD (Fig. 4-1B).
To validate our RNA-seq results, we used qRT-PCR to analyze the effects of
CsrA on fabB and ftnA expression, which seem to be regulated by CsrA at the level of
their transcription (Esquerré et al., 2016) (Potts et al., unpublished). As Fig. 4-3F shown,
csrA mutation significantly increased expression of fabB and ftnA, and csrA
overexpression dramatically decreased their expression, both in the csrD wild-type and
csrD mutant backgrounds. These results also support a CsrD-independent role of CsrA
on gene expression.
The finding that CsrA regulates gene expression largely independently of CsrD
does not support Model 2 in which CsrA mediates transcriptional changes primarily
through CsrD (Fig. 4-1B). This raises a question as to how CsrA affects the
transcriptional landscape. We suspect that it does this directly through its
posttranscriptional regulation of RNA stability and transcription elongation, and also
indirectly by controlling other transcriptional and posttranscriptional regulators. Indeed,
our data showed that CsrA affected the expression of 79 transcription regulators,
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including two component systems (TCS), σ factors, transcription factors, and enzymes
involved in the metabolism of the secondary messenger c-di-GMP (Table B-4). This is
consistent with recent studies in our group, which demonstrated that CsrA interacts with
many mRNAs encoding transcriptional regulators in vivo as determined using UV-
crosslinking immunoprecipitation and sequencing (CLIP-seq) (Potts et al., unpublished).
In addition, CsrA affected the abundance of 8 sRNAs in addition to CsrB and CsrC
(Table B-5). sRNAs have recently been discovered to participate in transcriptional
regulation via various transcriptional regulators (Göpel & Görke, 2012, Mika & Hengge,
2014, Lee & Gottesman, 2016, Mandin & Guillier, 2013). It seems that integration of
posttranscriptional regulation into transcriptional regulatory networks is a common
theme used by bacterial cells. As result, posttranscriptional regulators (such as CsrA
and sRNAs) can have vast effects on gene expression and control a broad range of
genes and cellular functions. Overall, our data suggest that rather than acting through
CsrD to alter transcript levels, CsrA mediates changes in the transcription of many
genes indirectly through other regulators.
CsrD Effects on Gene Expression Require CsrA
The next question we addressed was whether CsrD affects gene expression in a
CsrA-dependent manner. We identified in total of 74 genes that were differentially
expressed between the wild-type and csrD mutant strains. These data suggest that
CsrD has a limited effect on transcriptome compared to CsrA under our experimental
condition (Figs. 4-4 and 4-3A). Importantly, deletion of csrD in the csrA mutant
background did not change expression of any genes with the exception of csrD and
csrB (csrAD – csrA, Fig. 4-4). This indicated that almost all of CsrD effects on gene
expression require CsrA. Furthermore, while ectopic expression of csrA caused vast
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transcriptional changes in the csrAD mutant background (csrAD pCsrA – csrAD, Fig. 4-
3B), ectopic expression of csrD in this strain did not result in any changes in gene
expression except of csrD and csrB (csrAD pCsrD – csrAD, Fig. 4-4). These results
confirm that CsrA functions downstream of CsrD to regulate gene expression.
CsrB/C Have Strong Effects on Gene Expression in the Absence of CsrD
CsrB/C affect gene expression indirectly through sequestration of CsrA (Liu et al., 1997,
Weilbacher et al., 2003, Vakulskas et al., 2015). According to Model 1 (Fig. 4-1A),
CsrB/C can impact the transcriptional landscape independently from CsrD. As shown in
Fig. 4-5, deletion of csrBC resulted in changes in expression of 40 genes in the wild-
type background (csrBC - WT) and 218 genes in the csrD mutant strain background
(csrBCD - csrD). This suggested that CsrB/C do not require CsrD in order to affect gene
expression. The more effective effects of the csrBC deletion in the csrD mutant than in
the wild-type background was likely due to higher level of CsrB in the csrD mutant,
which reduces CsrA activity more than in the wild type strain and results in greater
changes to the CsrA regulon upon CsrB/C deletion. Furthermore, overexpression of
CsrB in the csrBCD mutant strain resulted in changes in expression of 912 genes (Fig.
4-5), suggesting that CsrB/C retain their global roles in regulating gene expression in
the absence of CsrD. These data together with our observation that CsrA globally
affects gene expression independently of CsrD (Fig. 4-3) demonstrated that neither
CsrB/C nor CsrA acts primarily through CsrD to mediate changes in gene expression, in
stark contrast to the prediction of Model 2 (Fig. 4-1B).
CsrD Regulates the Majority of its Target Genes in a CsrB/C Dependent Manner
Model 1 predicts that CsrD indirectly modulates CsrA activity through its effect on
CsrB/C levels (Suzuki et al., 2006). To test if CsrD depends on CsrB/C to mediate
95
changes in gene expression, we compared the impact of CsrD on the transcriptional
landscape in the presence and absence of CsrB/C. Differential expression analysis
revealed 60 genes that respond to csrD deletion in the wild-type background (csrD –
WT, Fig. 4-6A) but not in csrBC mutant background (csrBCD – csrBC, Fig. 4-6A),
suggesting that most of CsrD effects on gene expression require the presence of
CsrB/C. We also validated the dependence of CsrD-mediated ftnB expression on csrA
and CsrB/C using qRT-PCR and confirmed that CsrD lost its regulation of ftnB
expression in csrA or csrBC mutant background (Fig. 4-6B).
In the absence of CsrB/C, 26 genes still respond to CsrD (csrBCD – csrBC, Fig.
4-6A), suggesting that an alternative pathway may allow CsrD to mediates changes
through some unknown factor(s) rather than CsrB/C. The sRNA McaS has been
observed to serve as antagonists of CsrA and inhibits CsrA activity when overexpressed
(Jørgensen et al., 2013, Sterzenbach et al., 2013). Also, other sRNA were recently
discovered to directly bind CsrA in vivo in Samonella and E. coli (Holmqvist et al.,
2016)(Potts et al., unpublished). We predict that CsrD might mediate some changes in
gene expression through effects on other transcripts in a CsrA-dependent manner, it
might also function independently of CsrA. In addition, we observed 12 genes that were
regulated by CsrD only in csrBC mutant background (csrBCD – csrBC, Fig. 4-6A) but
not in wild-type background (csrD – WT, Fig. 4-6A). This implied that the CsrB/C
independent pathway might have opposite roles as the CsrB/C dependent pathway in
controlling expression of these 12 genes. Surprisingly, the expression level changes of
these 12 genes caused by csrD deletion in csrBC mutant background were not
complemented by ectopic expression of csrD (csrBCD pCsrD – csrBCD, Fig. 4-6A). The
96
reason for this requires further investigation. Taken together, these results strongly
support Model 1 (Fig. 4-1A) in which most of CsrD effects on CsrA activity are mediated
through its effect on CsrB/C turnover. However, CsrD might also regulate other RNAs
that affect CsrA activity. We cannot rule out the possibility that CsrD regulates a limited
number of genes independently of CsrA, but this possibility needs to be further
explored.
Conclusion
Overall, we provide evidence on a genome-wild scale that CsrA protein and
CsrB/C sRNAs mediate vast transcriptional changes independently of CsrD. Moreover,
CsrD primarily affects gene expression through its effects on CsrB/C and CsrA. In
addition, some of our data suggest that CsrD may affect expression of genes through
additional factors, something that warrants further investigation. Altogether, the use of
Epi-seq to clarify the epistatic relationships among the components of the Csr system
supports the original model on CsrD function, in which CsrA acts as the most
downstream regulator of gene expression in the Csr system (Model 1, Fig. 4-1A) and
our data do not support Model 2, wherein CsrD acts as the primary regulator of the
indirect effects of CsrA (Fig. 4-1B).
97
Figure 4-1. Two proposed model depicting the epistatic relationship between CsrA and CsrD in regulating mRNA abundance. A) CsrD affects gene expression through changes in CsrB/C levels, which then affect the activity of CsrA to directly or indirectly control mRNA expression (Suzuki et al., 2006). B) CsrA acts as a posttranscriptional regulator affecting RNA stability, but it mediates most of its indirect effects on mRNA abundance though transcriptional effects controlled by CsrD (Esquerré et al., 2016).
98
Figure 4-2. Properties of bacterial strains used in this study. A) Growth curve of the wild-
type and mutant strains carrying plasmids pBR322, pCsrA, pCsrD or pCsrB growing in KB medium at 37 °C. Control strains for pCsrA, pCsrD or pCsrB containing strains were transformed with plasmid pBR322. B) Northern blots of CsrB and CsrC RNAs in 37°C cultures of wild-type and mutant strains carrying plasmids pBR322, pCsrA, pCsrD, or pCsrB in KB medium at the transition from exponential to stationary phase of growth. RNA levels in mutant strains relative to those in the wild-type (WT) strain were shown at the bottom. ND, not detected. C, D and F) qRT-PCR analysis of the transcript levels of csrB, csrC and glgC genes in mutant strains with plasmids pBR322, pCsrA, pCsrD or pCsrB relative to those in the WT strain at the transition from exponential to stationary phase of growth. Error bars represent the mean results ± S.D. from three biological replicates. Asterisks indicate level of significance (*p≤0.05, **p≤0.01, ***p≤0.001) by Student’s t-test. E) Glycogen production in WT and mutant strains with plasmids pBR322, pCsrA, pCsrD or pCsrB were determined by iodine staining.
CsrC
WT
csrA
csrA
pCsr
A
csrD
csrD
pCsr
D
csrA
D
csrA
D p
Csr
A
csrA
D p
Csr
D
csrBC
csrB
CD
csrB
CD p
Csr
B
csrB
CD p
Csr
D
0
2
4
6
Re
lati
ve
exp
res
sio
n le
velCsrB
0
2
4
6
8
10
Re
lati
ve
ex
pre
ss
ion
le
ve
l
A B
C D
CsrB
1.0 0.1 1.3 ND 0.3 1.8 0.1 ND 1.3 ND
CsrC
1.0 0.1 0.8 ND 0.1 2.3 0.1 ND ND ND
csrB
C
cs
rAD
csrA
D
pCsr
A
csrA
D
pCsr
D
csrB
CD
csrB
CD
pC
srB
csrB
CD
pc
srD
Ratio to WT
WT csrA csrD
Ratio to WT
0 2 4 6 8 10 12 14 16 18 20 22 240.01
0.1
1
10
Time (Hour)
OD
600
WT
csrA
csrD
csrBC
csrAD
csrAD pCsrA
csrAD pCsrD
csrBCD
csrBCD pCsrB
csrBCD pCsrD
WT
csrA
csrA
pCsr
A
csrD
csrD
pCsr
D
csrA
D
csrA
D p
Csr
A
csrA
D p
Csr
D
csrB
C
csrB
CD
csrB
CD p
Csr
B
csrB
CD p
Csr
D
* *
* *
99
Figure 4-2. Continued.
100
Figure 4-3. CsrA retains its global role in regulating mRNA levels in the absence of
CsrD. A) Volcano Plot of the log2 fold change of RNA levels between the csrA
mutant and its isogenic wild-type strain. P-value ≤ 0.05 and a log2 fold
change higher than 1 or lower than − 1 were used as cutoff. The numbers of downregulated and upregulated genes (black dots) were shown on the top. B) Venn diagram depicting overlap of differentially expressed genes in csrA versus WT, csrAD versus csrD, and csrAD pCsrA versus csrAD. C, D and E) The log2 transformed fold change in RNA abundance caused by loss or overexpression of CsrA in the WT and/or csrD mutant backgrounds. Blue and red dots represent the genes that are only differentially expressed in one strain background. Black and grey dots represent the genes that are differentially expressed in both of the backgrounds, or neither of the background, respectively. The Spearman's correlation coefficients (ρ) were shown. F) qRT-PCR analysis of the transcript levels of acnA and ftnB genes in mutant strains csrA, csrD and csrAD carrying plasmids pBR322, pCsrA or pCsrD relative to those in the WT strain at the transition from exponential to stationary phase of growth. Error bars represent the mean results ± S.D. from three biological replicates. Asterisks indicate level of significance (*p≤0.05, **p≤0.01, ***p≤0.001) and n.s. indicates not significant by Student’s t-test.
101
Figure 4-3. Continued.
102
Figure 4-4. Venn diagram depicting the overlap of differentially expressed genes
induced by csrD deletion/overexpression in csrA WT and csrA mutant backgrounds. Specifically, differentially expressed genes in csrD versus WT, csrAD versus csrA, csrAD pCsrD versus csrAD were compared.
Figure 4-5. Venn diagram depicting the overlap of differentially expressed genes
induced by csrB/C deletion or csrB overexpression in csrD WT and csrD mutant backgrounds. Specifically, differentially expressed genes in csrBC versus WT, csrBCD versus csrD, csrBCD pCsrB versus csrBCD were compared.
103
Figure 4-6. CsrD regulate the majority of its target genes in a CsrB/C dependent
manner. A) Venn diagram depicting the overlap of differentially expressed genes in csrD versus WT, csrBCD versus csrBC, and csrBCD pCsrD versus csrBCD. B) qRT-PCR analysis of the transcript levels of ftnB gene in mutant strains carrying plasmids pBR322, pCsrA, pCsrD or pCsrB relative to those in the wild-type strain at the transition from exponential to stationary phase of growth. Error bars represent the mean results ± S.D. from three biological replicates. Asterisks indicate level of significance (*p≤0.05, **p≤0.01, ***p≤0.001) and n.s. indicates not significant by Student’s t-test.
104
CHAPTER 5 GENERAL DISCUSSION AND FUTURE PERSPECTIVES
Csr/Rsm sRNAs (for example E. coli CsrB/C) are non-coding sRNAs that
indirectly control gene expression by binding to and sequestering the activity of the
posttranscriptional regulatory proteins, CsrA/RsmA (Romeo et al., 2013, Vakulskas et
al., 2015). Synthesis and turnover of CsrB/C in E. coli are both controlled by
environmental signals and play important roles in governing CsrA activity and bacterial
lifestyle (Chavez et al., 2010, Huang et al., 2008, Edwards et al., 2011, Camacho et al.,
2015, Pannuri et al., 2016, Leng et al., 2016). The turnover of CsrB/C is initiated by
RNase E cleavage and subsequently faciliated by PNPase (Suzuki et al., 2006,
Vakulskas et al., 2016). This RNase E-dependent turnover of CsrB/C also requires
CsrD protein. The absence of CsrD significantly stabilized CsrB/C (Suzuki et al., 2006).
In this study, the physiological role of CsrD, the exact molecular mechanism how CsrD
facilitates CsrB/C turnover and whether CsrD has broader regulatory roles in addition to
regulating CsrB/C turnover were explored.
First, we revealed a physiological role of CsrD in coupling CsrB/C decay to
availability of preferred carbon sources. The CsrD effect is achieved by a direct
interaction of EIIAGlc of the glucose-specific PTS system to the EAL domain of CsrD. We
demonstrated that EIIAGlc regulates CsrB/C turnover in a phosphorylation dependent
manner and only the unphosphorylated form of EIIAGlc bound to CsrD and was capable
of activating CsrB/C turnover. On the other hand, the phosphorylated form of EIIAGlc
indirectly and modestly represses CsrB turnover via cAMP-Crp, reinforcing the positive
effect of unphosphorylated EIIAGlc on CsrB decay. This regulatory pathway couples
CsrB/C sRNA decay to the availability of a preferred carbon source, glucose. While
105
CsrB/C in strains lacking CsrD protein are extremely stable, the presence of CsrD in E.
coli ensures the high decay rates of CsrB/C in the presence of preferred carbon source
and poises the Csr system for rapid response to environmental stimuli.
This study also uncovered an important new way in which EIIAGlc shapes global
regulatory circuitry in response to nutritional status. Previous studies have identified
regulatory roles of EIIAGlc in carbon metabolism (Deutscher et al., 2014, Deutscher et
al., 2006), chemotaxis (Neumann et al., 2012), respiration/fermentation (Koo et al.,
2004), biofilm formation (Pickering et al., 2012) and virulence (Kim et al., 2010, Mazé et
al., 2014). Here, we showed that EIIAGlc facilitates the turnover of sRNAs CsrB/C in the
presence of glucose, resulting in decreased CsrB/C levels and therefore increased CsrA
activity to stimulate expression of genes for rapid growth.
In addition, this study uncovered a novel function of the EAL domain. A few
catalytically inactive EAL domains have been discovered to perform regulatory roles via
protein-protein interactions (Römling et al., 2013, Li et al., 2012, Tschowri et al., 2009,
Guzzo et al., 2009), but they all act as sensors to transmit information to another
protein. Here, the EAL domain of CsrD was revealed to act as a receiver and detect
signaling information from a sensory protein EIIAGlc. Also, it would be very intriguing to
figure out the molecular mechanism of the interaction between EAL domain and EIIAGlc
by crystallography.
More importantly, these findings revealed a new physiological influence on the
workings of the Csr system. Here we discovered that the presence of glucose
stimulates CsrB/C turnover and reduces CsrB/C levels. Previous studies have shown
that carboxylic acid-containing end products of carbon metabolism, such as acetate and
106
formate, stimulate CsrB transcription via the BarA-UvrY TCS (Chavez et al., 2010).
Thus, carbon availability is capable to mediate reinforcing effects on the levels of these
sRNAs through their synthesis pathway and the newly discovered turnover pathway.
This allows bacterial cells to rapidly alter the concentration of CsrA and properly
mediate gene expression in order to switch between active growth and stress resistant
growth upon changes in carbon availability. Of particular interest, even though CsrB/C
turnover rates are severely inhibited by the lack of EIIAGlc or CsrD, their levels are only
modestly changed due to feedback loops. We wonder that whether the turnover
pathway has a bigger impact in earlier stage upon changes in carbon availability and
the impact lessens over time when the transcriptional response takes over. It would be
of great interest to characterize the signaling dynamics of each pathway under different
carbon conditions, specifically, the real-time response patterns of CsrB/C turnover and
the transcriptional response could be investigated.
Besides, another question worthy of further investigation is whether other
environmental signals in addition to carbon availability affect CsrB/C turnover. The Csr
system captures environmental stimuli mainly through fluctuations of the steady state
levels of CsrB/C and then converges this into global regulation through CsrA. Multiple
regulatory factors that mediate stress responses, including stringent response
components (p)ppGpp/DksA (Edwards et al., 2011), the global stress σ factors RpoS
(Yakhnin et al., 2011), RNA chaperon Hfq (Suzuki et al., 2006), RNA helicases
Dead/SrmB (Vakulskas et al., 2014), and Crp (Pannuri et al., 2016) have been
discovered to control the transcription of CsrB/C. It would be interesting to know
107
whether these factors also regulate the Csr system through affecting the turnover of
CsrB/C.
Our recent work demonstrated that RNase E-mediated turnover of CsrB is
antagonistically controlled by CsrA and CsrD in E. coli (Leng et al., 2016). The binding
of CsrA to CsrB blocks the RNase E-mediated cleavage of CsrB both in vitro and in
vivo, while CsrD facilitates CsrB turnover by overcoming the CsrA-mediated protection
in vivo. In this study, we showed that although CsrD binds to CsrB in vitro, it did not
seem to bind CsrB in vivo and it failed to facilitate the cleavage of CsrA-protected CsrB
by RNase E in vitro. These data suggest that rather than functioning as an RNA binding
protein, CsrD might act indirectly and require other unknown factor(s) for its regulatory
role on CsrB/C turnover.
Proteins that are associated with CsrB in vivo were analyzed in this study by
immunoprecipitation and mass-spectrometry. In addition to CsrA, components of the
RNA degradosome, ribosomal proteins and PAP I were identified. While enolase was
strongly enriched along with purified CsrB, it does not influence CsrB/C turnover.
Whether other proteins, such as ribosomal proteins, are functionally relevant to CsrB
decay or have other functions remains to be further explored. Identifying the missing
factors involved in CsrB turnover will not only help to elucidate how CsrD works but also
provides new insights into the complexity of sRNAs degradation pathway. Genetic
screening can be conduced in future work to search for these factors.
In this study, we clarified the epistatic relationships of CsrA, CsrB/C and CsrD in
Csr system using Epi-seq. According to our previous study (Suzuki et al., 2006), CsrD
was proposed to affect gene expression through its effects on CsrB/C turnover and then
108
CsrA activity (Model 1, Fig. 4-1A). However, a recent transcriptomics study has
proposed an alternative model that CsrD works downstream of CsrA for regulating gene
expression and it is responsible for most of the indirect effects of CsrA on RNA
abundance (Model 2, Fig. 4-1B) (Esquerré et al., 2016). Our data illustrated on a
genome-wide scale that CsrD mediates changes in gene expression primarily through
its effects on CsrB/C and CsrA. Moreover, CsrA does not require CsrD to exert its
global effects on transcript levels, and its effects on indirect targets might be mediated
via other transcriptional and posttranscriptional regulators. In addition, our data
suggested that CsrD affects expression of some genes through an alternative pathway
independent of CsrB/C, suggestive of other regulatory role(s) of CsrD in addition to
facilitating CsrB/C turnover. Further investigation will be required to identify these
additional factors and pathways.
109
APPENDIX A SUPPLEMENTARY FIGURES
Figure A-1. EIIAGlc stimulates CsrB decay in the MG1655 csrDFLAG strain. Northern blots
depicting the effect of EIIAGlc on CsrB decay in strains MG1655 csrDFLAG and MG1655 Δcrr csrDFLAG. Half-lives were determined as in Figure 2-1.
Figure A-2. Decay of CsrB/C in a strain expressing EIIAGlc H91D is similar to that of EIIAGlc H91A. Decay rates of CsrB/C were determined in the Δcrr strain containing plasmid pCRRH91D, as described for the pCRRH91A-containing strain in Figure 2-1A and C.
Figure A-3. CRP has minimal or no effect on CsrD protein levels. Western blots depicting effects of crp deletion on CsrD protein levels. RpoB was used as loading control. CsrD protein levels in Δcrp relative to those in the wild-type strain (WT) were given. Standard derivations from triplicate experiments are indicated.
0.1 0.5 1.0 2.0
WT Δcrp WT Δcrp WT Δcrp WT Δcrp
OD600
1.5 ± 0.3 1.3 ± 0.1 1.0 ± 0.2 1.0 ± 0.1 Δcrp/WT
CsrDFLAG
RpoB
110
APPENDIX B SUPPLEMENTARY TABLES
Table B-1. Bacterial Strains
Strain Description Reference
MG1655 Prototrophic E. coli K12 Michael Cashel MG1655 Δcrr MG1655 with unmarked crr deletion This study (Chapter 2) MG1655 ΔcsrD MG1655 with unmarked csrD
deletion This study (Chapter 2)
MG1655 Δcrp MG1655 with marked crp disruption-Camr
This study (Chapter 2)
MG1655 ΔcyaA MG1655 with marked cyaA deletion-Kanr
This study (Chapter 2)
MG1655 Δcrr ΔcsrD
MG1655 with unmarked crr deletion and marked csrD deletion-Kanr
This study (Chapter 2)
MG1655 Δcrr Δcrp
MG1655 with unmarked crr deletion and marked crp disruption-Camr
This study (Chapter 2)
MG1655 Δcrr ΔcyaA
MG1655 with unmarked crr deletion and marked cyaA deletion-Kanr
This study (Chapter 2)
MG1655 csrDFLAG
MG1655 with in-frame, CTD 3X-FLAG tag at native csrD locus
This study (Chapter 2)
MG1655Δcrr csrDFLAG
csrDFLAG allele introduced by transduction using P1vir - Kanr
This study (Chapter 2)
MG1655 Δcrp csrDFLAG
csrDFLAG allele introduced by transduction using P1vir - Kanr
This study (Chapter 2)
MG1655 crrFLAG MG1655 with in-frame, CTD 3X-FLAG tag at native crr locus
This study (Chapter 2)
MG1655 H91A MG1655 with a His91Ala exchange (CAC to GCC) at native crr locus
This study (Chapter 2)
MG1655 H91A crrFLAG
crrFLAG allele introduced by transduction using P1vir
This study (Chapter 2)
BL21(DE3) Host for expression (Studier & Moffatt, 1986) DE5α Host for plasmid amplification (Woodcock et al., 1989) PW1096 Vibrio cholerae C6706str2 (Thelin & Taylor, 1996) PW1197 PW1096 with in frame mshH deletion This study (Chapter 2) PW1198 PW1096 with in frame crr deletion This study (Chapter 2) PW1207 PW1096 with in fram mshH and crr
deletions This study (Chapter 2)
MG1655 rne-1 ΔcsrB pCsrB
MG1655 rne-1 with marked csrB deletion-Gmr, harboring plasmid pCsrB
This study (Chapter 3)
MG1655 rne-1 ΔcsrB p2SLStrepto-CsrB
MG1655 rne-1 with marked csrB deletion-Gmr, harboring plasmid p2SLStrepto-CsrB
This study (Chapter 3)
111
Table B-1. Continued.
Strain Description Reference
MG1655 rne-1 ΔcsrB ΔcsrD pCsrB
MG1655 rne-1 ΔcsrB pCsrB with marked csrD deletion-Kanr
This study (Chapter 3)
MG1655 rne-1 ΔcsrB ΔcsrD p2SLStrepto-CsrB
MG1655 rne-1 ΔcsrB p2SLStrepto-CsrB with marked csrD deletion-Kanr
This study (Chapter 3)
MG1655 rne-1 ΔcsrB csrDFLAG pCsrB
MG1655 rne-1 ΔcsrB pCsrB with in-frame CTD 3X-FLAG tag at native csrD locus
This study (Chapter 3)
MG1655 rne-1 ΔcsrB csrDFLAG p2SLStrepto-CsrB
MG1655 rne-1 ΔcsrB p2SLStrepto-CsrB with in-frame CTD 3X-FLAG tag at native csrD locus
This study (Chapter 3)
MG1655 ΔcsrD pBYH4 MG1655 with marked csrD deletion-Kanr, harboring plasmid pBYH4
This study (Chapter 3)
MG1655 ΔcsrD p3FLAG-CsrD
MG1655 with marked csrD deletion-Kanr, harboring plasmid p3FLAG-CsrD
This study (Chapter 3)
XWC880 MG1655 with marked pgaC disruption-Camr
(Wang et al., 2004)
MG1655 csrA::gm MG1655 with csrA disrupted after amino acid position 50 - Gmr
(Vakulskas et al., 2016)
KDMG MG1655 with marked csrD deletion-Kanr
(Suzuki et al., 2006)
BW25113 csrB::kan
BW25113 with marked csrB deletion-Kanr
Baba, T 2006 (Baba et al., 2006)
BW25113 csrC::kan BW25113 with marked csrC deletion-Kanr
Baba, T 2006 (Baba et al., 2006)
Wild-type (WT)
MG1655 with marked pgaC disruption-Camr, carrying plasmid pBR322
This study (Chapter 4)
csrA
MG1655 with marked pgaC disruption-Camr and CsrA disruption-Camr, carrying plasmid pBR322
This study (Chapter 4)
csrD
MG1655 with marked pgaC disruption-Camr and csrD deletion-Kanr, carrying plasmid pBR322
This study (Chapter 4)
csrBC
MG1655 with marked pgaC disruption-Camr and unmarked csrB and csrC deletions, carrying plasmid pBR322
This study (Chapter 4)
112
Table B-1. Continued.
Strain Description Reference
csrAD
MG1655 with marked pgaC disruption-Camr, marked csrA disrutpion-Gmr and marked csrD deletion-Kanr, carrying plasmid pBR322
This study (Chapter 4)
csrA pCsrA
MG1655 with marked pgaC disruption-Camr and marked csrA disruption-Gmr, carrying plasmid pCsrA
This study (Chapter 4)
csrD pCsrD
MG1655 with marked pgaC disruption-Camr and marked csrD deletion-Kanr, carrying plasmid pCsrD
This study (Chapter 4)
csrAD pCsrD
MG1655 with marked pgaC disruption-Camr, marked csrA disrutpion-Gmr and marked csrD deletion-Kanr, carrying plasmid pCsrD
This study (Chapter 4)
csrAD pCsrA
MG1655 with marked pgaC disruption-Camr, marked csrA disrutpion-Gmr, and marked csrD deletion-Kanr, carrying plasmid pCsrA
This study (Chapter 4)
csrBCD
MG1655 with marked pgaC disruption-Camr, marked csrD deletion-Kanr and unmarked csrB and csrC deletions, carrying plasmid pBR322
This study (Chapter 4)
csrBCD pCsrB
MG1655 with marked pgaC disruption-Camr, marked csrD deletion-Kanr and unmarked csrB and csrC deletions, carrying plasmid pCsrB
This study (Chapter 4)
csrBCD pCsrD
MG1655 with marked pgaC disruption-Camr, marked csrD deletion-Kanr and unmarked csrB and csrC deletions, carrying plasmid pCsrD
This study (Chapter 4)
113
Table B-2. Plasmids used in this study.
Name Description Relevant Primers
Reference
pBR322 Cloning vector - Ampr Tetr N/A (Bolivar et al., 1977)
pCRR
pBR322 derivative carrying crr gene between EcoRI and HindIII sites- Ampr
P1/P2 This study (Chapter 2)
pCRRH91A pCRR derivative carrying crr with a His91Ala exchange (CAC to GCC)
P1/P17/P2
This study (Chapter 2)
pCRRH91D pCRR derivative carrying crr with a His91Asp exchange (CAC to GAC)
P1/P19/P2
This study (Chapter 2)
pBYH4
pBR322 derivative carrying csrD gene in EcoRI site- Ampr
N/A
(Suzuki et al., 2006)
pET24CRR pET24a derivative carrying crr gene between NdeI and XhoI sites
P3 /P4 This study (Chapter 2)
pDTM
pMAL-c5x derivative carrying DNA encoding 156-646 aa of CsrD between NcoI and EcoRI sites
P5/P10
This study (Chapter 2)
pDEAL
pMAL-c5x derivative carrying DNA encoding 156-385 aa of CsrD between NcoI and EcoRI sites
P6/P7
This study (Chapter 2)
pDGGDEF pMAL-c5x derivative carrying DNA encoding 156-223 and 393-646 aa of CsrD between NcoI and EcoRI sites
P6/P8 and P9/P10
This study (Chapter 2)
pDHAMP pMAL-c5x derivative carrying DNA encoding 192-646 aa of CsrD between NcoI and EcoRI sites
P11/P10
This study (Chapter 2)
pDcoil pMAL-c5x derivative carrying DNA encoding 156-199 and 220-646 aa of CsrD between NcoI and EcoRI sites
P6/P12 and P13/P10
This study (Chapter 2)
pEAL pMAL-c5x derivative carrying DNA encoding 393-646 aa of CsrD between NcoI and EcoRI sites
P14/P10
This study (Chapter 2)
pKOV Vector for homologous recombination
N/A (Link et al., 1997)
pKOVH91A pKOV derivative carrying crr with a His91Ala exchange (CAC to GCC) between NotI and BamHI sites
P15/P17and P18/ P16
This study (Chapter 2)
pCsrB pBR322 derivative carrying csrB gene with 494bp upstream and 36bp downstream between BamHI and HindIII sites- Ampr
csrB-494upBamHI/csrB-36downHindIII
This study (Chapters 3 and 4)
114
Table B-2. Continued.
Name Description Relevant Primers
Reference
p2SLStrepto-CsrB
pBR322 derivative expressing Strepto-CsrB- Ampr
p2SLStrepto-CsrB-F/R
This study (Chapter 3)
p3FLAG-CsrD pBR322 derivative expressing 3FLAG tagged CsrD- Ampr
p3FLAG-CsrD-F/R
This study (Chapter 3)
p2VR112 csrA gene cloned into the EcoRI-BamHI sites of pBR322 - Ampr
N/A (Vakulskas et al., 2016)
115
Table B-3. Primers used in this study.
Primer Sequence (5’-3’) Function
P1 CAGTACCAGGAATTCTTTACACTTTATGCTTCCGGCTCGTATATTGTGTGGAAGAAATAATTTTGTTTAACTTTAAG
pCRR and pH91D construction
P2 CAGGACCATAAGCTTTTACTTCTTGATGCGGATAACCGGGGT
pCRR construction
P3 ACATGATCTCATATGGGTTTGTTCGATAAACTGAAATCTC
pET24CRR construction
P4 ACATGATCTCTCGAGCTTCTTGATGCGGATAACCGGGGTT
pET24CRR construction
P5 CCGATTCCATGGGCCGCTGGTTACAACGGCAACTTGCCG
pDTM construction
P6 CATGCCATGGGCCGCTGGTTACAACGGCAACTTGC
pDEAL, pDGGDEF and pDcoil construction
P7 ACCGGAATTCTTAGTAAATAGCCCAGCTATTGCCGC
pDEAL construction
P8 AACATTACCGCGTCCATAAGAGCGGATCAG pDGGDEF construction
P9 CTGATCCGCTCTTATGGACGCGGTAATGTT pDGGDEF construction
P10 ACCGGAATTCTTAAACCGAGTATCTTTGTGAATAT
pDTM, pDGGDEF, pDHAMP, pDcoil and pEAL construction
P11 CATGCCATGGGCCCGCCCAGAACCAGCAGTGC pDHAMP construction
P12 GGCGGCATAAGAGCGGATCAGCGCACTGCTGGTTCT
pDcoil construction
P13 AGAACCAGCAGTGCGCTGATCCGCTCTTATGCCGCC
pDcoil construction
P14 CATGCCATGGGCGGACGCGGTAATGTTCGCTGGCGTA
pEAL construction
P15 ATAAGAATGCGGCCGCAAAGATCTGCCAGCTATTACGCTGG
pKOVH91A construction
P16 CGCGGATCCTGATAGCCGATTTGACTGCCAGAAT pKOVH91A construction
P17 GGTGTCGATACCGAAGGCGACGAACAGTTCAAC pCRRH91A and pKOVH91A construction
P18 GTTGAACTGTTCGTCGCCTTCGGTATCGACACC pKOVH91A construction
P19 GGTGTCGATACCGAAGTCGACGAACAGTTCAAC pCRRH91D construction
116
Table B-3. Continued.
Primer Sequence (5’-3’) Function
S1 CGTAACCGTGGGTGAAACCCCGGTTATCCGCATCAAGAAGGACTACAAAGACCATGACGG
MG1655 crrFLAG
construction S2 AAATGGCGCCGATGGGCGCCATTTTTCACTGCG
GCAAGAACATATGAATATCCTCCTTAG MG1655 crrFLAG
construction S3 TGATACTAACGTGAAAAAATATTCACAAAGATAC
TCGGTTGACTACAAAGACCATGACGG MG1655 csrDFLAG construction
S4 AGCGCGCATTATTCTACGTGAAAACGGATTAAACGGCAGGCATATGAATATCCTCCTTAG
MG1655 CsrDFLAG construction
R1 GTAATACGACTCACTATAGTCGACAGGGAGTCAGACAAC
RNA CsrB synthesis
R2 AAAAAAAGGGAGCACTGTATTCACAGCGCTCCCGGTTCGTTTCGCAG
RNA CsrB synthesis
R3 GTAATACGACTCACTATAGGATAGAGCGAGGACGCTAACAGGAAC
RNA CsrC synthesis
R4 AAGAAAAAAGGCGACAGATTACTCTGTCGCCTTTTTTCCTGACTC
RNA CsrC synthesis
R5 GTAATACGACTCACTATAGGCCTTTGAATTGTCCATATAGAACAC
RNA rpsT synthesis
R6 AAAAAAACCCGCTTGCGCGGGCTTTTTCACAAAGCTTCAGC
RNA rpsT synthesis
R7 TAATACGACTCACTATAGGGTAGATGCTCATTCCATCTC
RNA GlmZ synthesis
R8 AAAAAAACGCCTGCTCTTATTACGGAGCAGGCGTTAAAAC
RNA GlmZ synthesis
RP1 TAATACGACTCACTATAGGGTAAAAGGTGCTCCCTGCATCTAATC
V. cholerae CsrB probe synthesis
RP2 TGGTGATCTTCAGGAAGAAGAATCG V. cholerae CsrB probe synthesis
RP3 TAATACGACTCACTATAGGGATCCTTTCAGCGAACTCCGAGCATC
V. cholerae CsrC probe synthesis
RP4 CAGGATGAGAAGTGGTGAGGATGAC V. cholerae CsrC probe synthesis
RP5 TAATACGACTCACTATAGGGCAATCCCGCTACTAATAGGTGCTCC
V. cholerae CsrD probe synthesis
RP6 CAAGGATTGGTCATCTTCAGGACGA V. cholerae CsrD probe synthesis
RP7 GTAATACGACTCACTATAGGTTCGTTTCGCAGCATTCCAG
E. coli CsrB probe synthesis
RP8 GCGTTAAAGGACACCTCCAGG E. coli CsrB probe synthesis
RP9 GTAATACGACTCACTATAGGTCTTACAATCCTTGCAGGC
E. coli CsrC probe synthesis
RP10 GAGGACGCTAACAGGAACAATG E. coli CsrC probe synthesis
117
Table B-3. Continued.
Primer Sequence (5’-3’) Function
3FLAG-CsrD-F
TGATATCGACTACAAAGATGACGACGATAAATAGTAACCTGCCGTTTAATCCGTTTTCA
p3FLAG-CsrD construction
3FLAG-CsrD-R
TGATCTTTATAATCACCGTCATGGTCTTTGTAGTCAACCGAGTATCTTTGTGAATATTTT
p3FLAG-CsrD construction
2SLStrepto-CsrB-F
GCAAGGGCACCACGGTCGGATCCCACTTCTGCAGGACACACCAGGAT
p2SLStrepto-CsrB construction
2SLStrepto-CsrB-R
GGGCAGAAGTCCAAATGCGATCCCACTTCGTTGTCTGACTCCCTGTCG
p2SLStrepto-CsrB construction
csrB-494upBamHI
CGCGGATCCATATGCACGCGCAGTTTGGCGATAT pCsrB construction
csrB-36downHindIII
CCCAAGCTTACCTCAATAAGAAAAACTGCCGCGA pCsrB construction
70upCsrA
TGGCGTTATATGATGGATAATGCCG csrA::gm confirmation
100downCsrA
GAGACTTAAGTTGAATGAACGGGAG csrA::gm confirmation
118
Table B-4. Transcription regulators regulated by CsrA
Gene Locus Gene
Regulator Type Directon Description
b2127 mlrA Transcription factor Repressed Transcriptional regulator of csgD
b3555 yiaG Transcription factor Repressed
Predicted transcriptional regulator, function unknown
b1306 pspC Transcription factor Repressed
Positive regulatory gene, cooperatively with PspB; facilitates binding of PspA to PspB; membrane protein; dimer
b0313 betI Transcription factor Repressed
Transcriptional repressor for the betIBA-betT divergent operon, choline-inducible
b2664 csiR Transcription factor Repressed Repressor of csiD promoter
b1399 paaX Transcription factor Repressed
Repressor for the paa operon, phenylacetyl-CoA induced
b4396 rob Transcription factor Repressed
Right oriC-binding transcriptional activator, AraC family
b1299 puuR Transcription factor Repressed
Repressor for the divergent puu operons, putrescine inducible; putrescine utilization pathway
b4135 yjdC Transcription factor Repressed
Putative transcriptional repressor, function unknown
b4313 fimE Transcription factor Repressed
Site-specific recombinase, fimA promoter inversion; biased towards the "ON to OFF" fimbriae phase switching direction
b2706 srlM Transcription factor Repressed
srl operon transcriptional activator, sorbitol-responsive
b2537 hcaR Transcription factor Repressed
Transcriptional activator for the hca operon; inducd by 3-phenylpropionate and cinnamic acid; autoregulatory
b3995 rsd Transcription factor Repressed
Regulates RNA polymerase holoenzyme formation; interacts with free σ 70 and core RNA polymerase; stationary phase protein; anti-σ
b1217 chaB Transcription factor Repressed
Accessory and regulatory protein for chaA
b3023 ygiV Transcription factor Repressed
Represses mcbR, involved in biofilm regulation
b0487 cueR Transcription factor Repressed
Activator of copper-responsive regulon genes cueO and copA; MerR homolog
b2398 yfeC Transcription factor Repressed
Predicted DNA-binding protein, DUF1323 family
119
Table B-4. Continued.
Gene Locus Gene
Regulator Type Directon Description
b0435 bolA Transcription factor Repressed
Stationary-phase morphogene, transcriptional repressor; predicted reductase; regulates mreB, dacA, dacC, and ampC transcription;
b4340 yjiR Transcription factor Repressed
Predicted HTH transcriptional regulator with aminotransferase domain, function unknown; MocR family
b2980 glcC Transcription factor Repressed
Transcriptional reperssor for glc operon, glycolate-binding
b0330 prpR Transcription factor Repressed
Transcriptional regulator of prp operon; propionate catabolism via 2-methylcitrate cycle, characterized primarily in Salmonella
b1422 ydcI Transcription factor Repressed Putative transcriptional regulator
b1014 putA Transcription factor Repressed
Proline dehydrogenase and repressor for the putAP divergon
b1384 feaR Transcription factor Repressed
Transcriptional activator for tynA and feaB, AraC family
b2427 murR Transcription factor Repressed
Repressor for murPQ, MurNAc 6-P inducible
b3604 lldR Transcription factor Repressed
Dual role activator/repressor for lldPRD operon
b1622 malY Transcription factor Repressed
Antagonist of MalT transcriptional activator of maltose regulon, binds MalT in absence of maltotriose; cysteine desulfhydrase
b0603 ybdO Transcription factor Repressed
Required for swarming phenotype, function unknown; probable LysR-family transcriptional regulator
b0020 nhaR Transcription factor Repressed
Positive regulator of nhaA, Na(+)-dependent
b0145 dksA Transcription factor Repressed
RNAP-binding protein modulating ppGpp and iNTP regulation; reduces open complex half-life on rRNA promoters; removes transcriptional roadblocks to replication
b3680 yidL Transcription factor Repressed
Predicted transcriptional regulator, AraC family, function unknown
b0064 araC Transcription factor Repressed
Transcriptional activator for the ara regulon
120
Table B-4. Continued.
Gene Locus Gene
Regulator Type Directon Description
b2399 yfeD Transcription factor Repressed
Predicted DNA-binding protein, DUF1323 family
b4365 yjjQ Transcription factor Repressed
Putative transcriptional regulator, function unknown; H-NS-repressed, dimeric
b3021 mqsA Transcription factor Repressed
Antitoxin for MqsR toxin; transcriptional repressor
b2846 yqeH Transcription factor Repressed
Predicted LuxR family transcriptional regulator; part of T3SS PAI ETT2 remnant
b1450 mcbR Transcription factor Repressed
MqsR-controlled colanic acid and biofilm regulator; represses mcbA
b0346 mhpR Transcription factor Repressed
Transcriptional activator, mhp operon; utilizes MHP
b1499 ydeO Transcription factor Repressed
UV-inducible global regulator, EvgA-, GadE-dependent; transcriptional activator for mdtEF; AraC family
b0076 leuO Transcription factor Repressed
Pleiotropic transcriptional regulator; regulates dsrA; relieves bgl silencing, multi-copy represses cadC
b2491 hyfR Transcription factor Repressed
Formate-sensing regulator for hyf operon
b3022 mqsR Transcription factor Repressed
GCU-specific mRNA interferase, toxin-antitoxin pair MqsRA; motility, quorum-sensing biofilm regulator;
b4366 bglJ Transcription factor Repressed
Transcriptional activator for the silent bgl operon; requires the bglJ4 allele to function; LuxR family
b2709 norR Transcription factor Repressed
Transcription regulator for norVW, NO-responsive; σ 54-dependent activator with a GAF domain
b0294 ecpR Transcription factor Repressed
Putative transcriptional regulator for the ecp operon
b2531 iscR Transcription factor Repressed
isc operon transcriptional repressor; suf operon transcriptional activator; icsR regulon regulator;
b2561 yfhH Transcription factor Repressed
putative DNA-binding transcriptional regulator
b0564 appY Transcription factor Activated
Global transcription regulator, AraC family, DLP12 prophage
b2714 ascG Transcription factor Activated
Repressor of asc operon; inducer unknown; prpBC operon repressor
121
Table B-4. Continued.
Gene Locus Gene
Regulator Type Directon Description
b3556 cspA Transcription factor Activated
Cold-inducible RNA chaperone and antiterminator; aids gene expression at low temperature;
b3261 fis Transcription factor Activated
Transcriptional activator for rRNA operons, bends DNA; interacts with RNAP; nucleoid-associated protein;
b1891 flhC Transcription factor Activated
Transcriptional activator of flagellar class II operons; forms heterotetramer with FlhD;
b1892 flhD Transcription factor Activated
Transcriptional activator of flagellar class II operons; forms heterotetramer with FlhC
b1658 purR Transcription factor Activated Purine regulon repressor
b3481 nikR Transcription factor Activated
Nickel-responsive regulator of the nik operon; homodimer
b4116 adiY Transcription factor Activated
Transcriptional activator for adiA, AraC family
b0450 glnK Transcription factor Activated
Potent activator of NRII (GlnL/NtrB) phosphatase; trimeric.
b4128 ghoS Transcription factor Activated
Antitoxin of GhoTS toxin-antitoxin pair; endonuclease for ghoT mRNA
b3071 yqjI Transcription factor Activated
Transcriptional repressor for yqjH, nickel- or iron-inducible; autorepressor
b0483 ybaQ Transcription factor Activated
Predicted transcriptional regulator, function unknown
b1983 yeeN Transcription factor Activated
UPF0082 family protein, function unknown
b1988 nac Transcription factor Activated
Repressor of gdhA transcription; RpoN, GlnG (NtrC) regulons; pleiotropic effects
b3869 glnL Sensor kinase Activated
Bifunctional kinase/phosphatase, nitrogen regulator II, NRII; homodimeric
b1222 narX Sensor kinase Repressed
Two-component nitrate/nitrite sensor-transmitter protein; NarL is cognate regulator;
b1609 rstB Sensor kinase Repressed
Sensory histidine kinase of RstAB two-component system, low Mg-responsive via PhoQP
b2078 baeS Sensor kinase Repressed
Sensor kinase for mdtABCD, acrD and spy
122
Table B-4. Continued.
Gene Locus Gene
Regulator Type Directon Description
b2503 yfgF Phosphodiesterase Activated
Cyclic-di-GMP phosphodiesterase, anaerobic; dual domain protein; defective cyclase domain; predicted membrane sensor protein
b1489 dosP Phosphodiesterase Repressed
Heme-regulated oxygen sensor, c-di-GMP phosphodiesterase; biofilm regulator;
b1815 yoaD Phosphodiesterase Repressed
Predicted membrane-anchored cyclic-di-GMP phosphodiesterase; regulation of cellulose production
b1285 gmr Phosphodiesterase Repressed
Cyclic-di-GMP phosphodiesterase; csgD regulator;
b1956 yedQ Diguanylate cyclase Repressed
Predicted membrane-anchored diguanylate cyclase
b2067 yegE Diguanylate cyclase Repressed
Predicted diguanylate cyclase, dual domain protein; defective phosphodiesterase domain;
b1341 ydaM Diguanylate cyclase Repressed
Diguanylate cyclase, csgD regulator; also regulates GGDEF protein YaiC (AdrA)
b1535 dgcZ Diguanylate cyclase Repressed Diguanylate cyclase, zinc-sensing
b1025 ycdT Diguanylate cyclase Repressed
Diguanylate cyclase, membrane-anchored
b1490 dosC Diguanylate cyclase Repressed
Diguanylate cyclase, binds oxygen, positive biofilm regulator; cold- and stationary phase-induced
b2741 rpoS σ factor Repressed RNA polymerase subunit, stress and stationary phase σ S; σ 38
b4293 fecI σ factor Repressed RNA polymerase σ-19 factor; fecA promoter RNAP σ factor
123
Table B-5. sRNAs regulated by CsrA at the transcript level
Gene Locus Gene Direction Description
b4458 oxyS Repressed OxyS sRNA activates genes that detoxify oxidative damage
b4577 sgrS Repressed sRNA that destabilzes ptsG mRNA
b4441 glmY Repressed sRNA activator of glmS mRNA, glmZ processing antagonist
b4444 omrA Repressed sRNA downregulating OM proteins and curli; positively regulated by OmpR/EnvZ
b4445 omrB Repressed sRNA downregulating OM proteins and curli; positively regulated by OmpR/EnvZ
b4459 ryjA Repressed Novel sRNA, function unknown
b4698 mgrR Repressed sRNA affecting sensitivity to antimicrobial peptides; regulated by PhoPQ and Mg2+
b4699 fnrS Activated FNR-activated anaerobic sRNA; mediates negative FNR regulation; Hfq-dependent
124
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BIOGRAPHICAL SKETCH
Yuanyuan Leng was born in Weifang, Shandong, China. She received her
bachelor’s degree in biotechonology from Shandong University, China in June of
2009. She then joined the program of State Key Laboratory of Microbial Technology at
Shandong University and received her master’s degree in microbiology in June of
2012. After that, in August of 2012, she joined the graduate program at the
Department of Microbiology and Cell Science at the University of Florida. After three
rotations, she joined Dr. Tony Romeo’s lab and worked with him to study the
regulation of the RNase E-mediated turnover of sRNAs CsrB/C. She received her
Ph.D. from the University of Florida in the summer of 2017. Yuanyuan Leng plans to
continue research training as a postdoctoral associate at National Cancer Institute.
