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Summer 2015

Clostridium difficile: shedding light on pathogenesis Clostridium difficile: shedding light on pathogenesis

Eric M. Ransom University of Iowa

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Copyright © 2015 Eric M. Ransom

This dissertation is available at Iowa Research Online: https://ir.uiowa.edu/etd/5828

Recommended Citation Recommended Citation Ransom, Eric M.. "Clostridium difficile: shedding light on pathogenesis." PhD (Doctor of Philosophy) thesis, University of Iowa, 2015. https://doi.org/10.17077/etd.x8v5w8yj

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CLOSTRIDIUM DIFFICILE: SHEDDING LIGHT ON PATHOGENESIS

by

Eric M. Ransom

A thesis submitted in partial fulfillment of the requirements for the Doctor of

Philosophy degree in Microbiology in the Graduate College of

The University of Iowa

August 2015

Thesis Supervisors: Associate Professor Craig Ellermeier Associate Professor David Weiss

Copyright by

ERIC M. RANSOM

2015

All Rights Reserved

Graduate College The University of Iowa

Iowa City, Iowa

CERTIFICATE OF APPROVAL

_______________________

PH.D. THESIS

_______________

This is to certify that the Ph.D. thesis of

Eric M. Ransom

has been approved by the Examining Committee for the thesis requirement for the Doctor of Philosophy degree in Microbiology at the August 2015 graduation.

Thesis Committee: _____________________________________ Craig Ellermeier, Thesis Supervisor

_____________________________________ David Weiss, Thesis Supervisor

_____________________________________ Patrick Schlievert

_____________________________________ Alexander Horswill

_____________________________________ Linda McCarter

_____________________________________ Kim Brogden

To my family, friends, & Gina.

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ACKNOWLEDGMENTS

I would like to acknowledge my co-mentors Craig Ellermeier and David Weiss. They are

true intellectuals who openly share their wealth of knowledge and excitement for the unknown.

I cannot thank them enough for their training, thoroughness, and patience. May the Brain Trust

live on!

I would like to thank past and current members of the Ellermeier and Weiss labs. In

particular, thank you Kyle Williams and Matthew Jorgenson who provided the ideal lab

environment, experimental advice, and unspeakable memories. Some stories are better left

untold. Also thank you to Jessica Hastie and Atsushi Yahashiri for your insight and lunch

conversations. A special thanks to “T” as well.

I have to thank my committee. Their advice was not only helpful but essential. If I ever

needed assistance or access to equipment, I could always count on them. I also wish to thank

other faculty, especially Linda Knudtson for the teaching experiences. Your influence will

forever be evident in my teaching. Also, thank you to Dr. Brad Ford and Dr. Dan Diekema for the

experiences in a clinical microbiology laboratory at the University of Iowa Hospitals and Clinics.

To my parents Dennis and Sandi, I want to thank you for the continuous support. Your

voices over the phone and visits to Iowa City always lessened the stress of graduate school. To

my brother Brian, sister-in-law Maria, and nephews Gavin and Preston, thank you for the

support, vacation-escapes, and understanding. To my Grandma, thank you for the calming

phone calls and please keep sending cookies after I graduate. I also must mention my late

grandparents from whom I learned my love of music and passion for education.

For their support and extracurricular activities, I would like to thank my fellow scholars

and friends: Justin “Petey” Peterson, Allen Neuharth, Nathan Chase, Jeremy Fischbach, my

goddaughter Zoie, Kevin Hofer, Lars Hovden, Katie Hassebroek, Kate Ahlers, Luke Watson, Justin

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Plasket, Carolyn Linke, Steven Ewart, Harty Boys (Jeff Nolz, Martin Richer, Bram Slutter), micro

softball team, volleyball team, and soccer team.

Lastly, I thank the essential Gina. None of this would be possible without you. You

listened, motivated, encouraged, assisted, edited, critiqued, celebrated, traveled, and loved.

Thank you.

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ABSTRACT

Clostridium difficile is a strictly anaerobic, spore-forming bacterium that is linked to over

250,000 infections annually in the United States. One of the greatest challenges facing C.

difficile research has been the lack of genetic tools. This limited repertoire is due, in part, to the

anaerobic nature of C. difficile. For example, most fluorescent protein reporters require O2 for

chromophore maturation. Here, we demonstrate that O2-dependent fluorescent proteins

produced anaerobically can acquire fluorescence after cells are fixed with cross-linkers to

preserve native patterns of protein localization. This was shown using the blue and the red

codon-optimized fluorescent proteins, CFPopt and mCherryOpt, respectively.

Little is known about cell division in C. difficile. Here we identify and characterize a

three-gene operon encoding cell division proteins found only in C. difficile and a small number of

closely related bacteria. These proteins were named MldA, MldB, and MldC, for midcell

localizing division proteins. MldA is predicted to be a membrane protein with coiled-coil

domains and a peptidoglycan-binding SPOR domain. MldB and MldC are predicted to be

cytoplasmic proteins; MldB has two predicted coiled-coil domains, while MldC lacks obvious

conserved domains or sequence motifs. Mutants of mldA or mldB had morphological defects,

including loss of rod shape (a curved cell phenotype) and inefficient separation of daughter cells

(a chaining phenotype). Fusions of CFPopt to MldA, MldB, and MldC revealed that all three

proteins localize sharply to the division site. Mutants lacking the Mld proteins are severely

attenuated for pathogenesis in a hamster model of C. difficile infection. Because all three Mld

proteins are essentially unique to C. difficile, they could be exploited as targets for antibiotics

that combat C. difficile without disrupting the intestinal microbiome.

C. difficile pathogenesis is mediated primarily by two large exotoxins called Toxin A

(TcdA) and Toxin B (TcdB). Transcription of tcdA and tcdB depends on TcdR, an alternative

v

sigma factor for RNA polymerase. Previous studies have shown both toxins are produced upon

entry into stationary phase, and that this response is mediated in part by the CodY repressor,

which senses GTP and branched chain amino acids. Here we used mCherryOpt as a reporter of

gene expression to visualize toxin expression at the level of individual cells. This approach led to

the unexpected discovery that only a subset of cells in the population induces expression of tcdA

(and tcdB under specific conditions). In other words, toxin production is a “bistable” phenotype.

Further experiments indicated TcdR plays a central role in mediating bistability, while CodY

makes a minor but still significant contribution to bistability. Why it is advantageous for only a

subset of C. difficile cells to produce toxin is not known, but one interesting possibility is related

to conflicting requirements for transmission to a new host. Some cells produce toxin to provoke

diarrhea while other cells differentiate into spores that can survive exposure to air.

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PUBLIC ABSTRACT

The bacterium Clostridium difficile has been identified by the Centers for Disease

Control and Prevention as an “urgent” threat to public health, the highest threat level. C.

difficile is linked to over 250,000 infections and 14,000 deaths annually in the United States.

While there is much interest in studying C. difficile, the field has been hindered by the lack of

genetic tools. This is due, in part, because C. difficile is a strict anaerobe and thus oxygen-

dependent tools cannot be utilized. Here, we discuss a new technique that uses oxygen-

dependent fluorescent proteins in a strictly anaerobic environment (i.e. C. difficile). Since

oxygen-dependent fluorescent proteins are powerful genetic tools, our findings enable

researchers to study C. difficile as never before. In addition, we optimized our technique in C.

difficile by developing a new red fluorescent protein. We have used these tools to study

localization of novel cell division proteins in C. difficile and the regulation of virulence gene

expression. Taken together, our work has led to novel insights into C. difficile pathogenesis that

could eventually lead to the development of new therapeutics.

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TABLE OF CONTENTS

LIST OF TABLES ................................................................................................................................. x LIST OF FIGURES ...............................................................................................................................xi

LIST OF ABBREVIATIONS ................................................................................................................ xiii

CHAPTER

I. INTRODUCTION ........................................................................................................... 1 History of Clostridium difficile (Bacillus difficilis) ........................................................ 1 Clostridium difficile...................................................................................................... 2 Epidemiology ............................................................................................................... 5 Clinical aspects of Clostridium difficile ........................................................................ 6 Clinical diagnosis ....................................................................................................... 10 Toxins of C. difficile ................................................................................................... 10 Pathogenicity Locus (PaLoc) ...................................................................................... 12 Regulation of toxin expression by metabolic inputs ................................................. 14 Overview of chapters ................................................................................................ 19

II. IDENTIFICATION AND CHARACTERIZATION OF A GENE CLUSTER REQUIRED FOR PROPER ROD SHAPE, CELL DIVISION, AND PATHOGENESIS IN CLOSTRIDIUM DIFFICILE ............................................................................................ 30 Introduction .............................................................................................................. 30 Materials and Methods ............................................................................................. 31

Strains, media, and growth conditions ............................................................. 31 Plasmid and strain construction ....................................................................... 32 Intergenic RT-PCR and qRT-PCR ....................................................................... 34 Morphology of mldA::erm and mldB::erm ....................................................... 34 Transmission electron microscopy ................................................................... 35 Localization of CFP-Mld fusion proteins in C. difficile ...................................... 35 Localization of FtsZ-GFP in E. coli ..................................................................... 36 Immunoblot analysis ........................................................................................ 36 Spore preparation............................................................................................. 38 Syrian hamster model of C. difficile infection................................................... 38

Results ....................................................................................................................... 39 Bioinformatic identification of a cell division gene cluster .............................. 39 The mldABC genes are co-transcribed ............................................................. 40 mld mutations impair cell division, cell elongation, and cell shape ................. 41 Complementation of insertions in mldA or mldB requires the entire mldABC operon ................................................................................................. 43 Overproduction of MldAB impairs cell division ................................................ 44 The Mld proteins localize to the midcell .......................................................... 45 mld mutants are attenuated for pathogenesis in a hamster model of C. difficile infection ............................................................................................... 47

Discussion .................................................................................................................. 48

III. USE OF MCHERRY RED FLUORESCENT PROTEIN FOR STUDIES OF PROTEIN LOCALIZATION AND GENE EXPRESSION IN CLOSTRIDIUM DIFFICILE ........................ 81

viii

Introduction .............................................................................................................. 81 Materials and Methods ............................................................................................. 82

Strains, media, and growth conditions ............................................................. 82 Plasmid and strain construction ....................................................................... 83 Bioinformatics ................................................................................................... 84 Comparison of mCherryOpt, CFPopt, and GFPmut2 ........................................ 84 Kinetics of fluorescence acquisition ................................................................. 85 Protein localization ........................................................................................... 85 Quantifying expression of PpdaV::mCherryOpt .................................................. 86 Fixation protocol ............................................................................................... 86 Microscopy ....................................................................................................... 87 Plasmid Copy Number ...................................................................................... 88 Plasmid Stability ............................................................................................... 88 Nucleotide sequence accession number .......................................................... 89

Results ....................................................................................................................... 89 mCherryOpt is superior to CFP in C. difficile .................................................... 89 Application of mCherryOpt to protein localization .......................................... 90 Application of mCherryOpt as a reporter of gene expression ......................... 92 Plasmid stability and copy number .................................................................. 93

Discussion .................................................................................................................. 94

IV. TOXIN GENE EXPRESSION IS BISTABLE IN C. DIFFICILE............................................ 122

Introduction ............................................................................................................ 122 Materials and Methods ........................................................................................... 124

Strains, media, and growth conditions ........................................................... 124 Plasmid and strain construction ..................................................................... 125 Fixation protocol ............................................................................................. 127 Microscopy ..................................................................................................... 127 Flow cytometry ............................................................................................... 128 Plate reader .................................................................................................... 128 RNA isolation from fixed cells ......................................................................... 128 qRT-PCR .......................................................................................................... 129

Results ..................................................................................................................... 129 tcdA expression is bistable ............................................................................. 129 Expression of tcdA is bistable in multiple C. difficile ribotypes ...................... 130 Glucose alters the bistable expression of tcdA .............................................. 131 Toxin bistability is dependent upon TcdR ...................................................... 131 Toxin bistability is not dependent upon CodY ................................................ 132 Preliminary evidence for bistable expression of tcdB .................................... 133

Discussion ................................................................................................................ 133

V. FUTURE DIRECTIONS ............................................................................................... 160 Chapter II ................................................................................................................. 160 Chapter III ................................................................................................................ 162 Chapter IV ............................................................................................................... 164

REFERENCES ................................................................................................................................. 168

ix

LIST OF TABLES

Table

2.1 Strains used in chapter two ................................................................................................. 52

2.2 Oligonucleotide primers used in chapter two ..................................................................... 54

2.3 Plasmids used in chapter two .............................................................................................. 56

2.4 Sequence identities and E-values of MldABC homologs ..................................................... 57

2.5 Chaining phenotypes of mld mutants .................................................................................. 58

2.6 Complementation of mld mutants ...................................................................................... 59

2.7 Dependency of localization on other Mld proteins ............................................................. 60

3.1 Strains used in chapter three ............................................................................................... 97

3.2 Oligonucleotide primers used in chapter three ................................................................... 98

3.3 Plasmids used in chapter three .......................................................................................... 100

3.4 Codon usage in mCherryOpt .............................................................................................. 101

3.5 Plasmid stability ................................................................................................................. 102

3.6 Plasmid copy number......................................................................................................... 103

4.1 Strains used in chapter four ............................................................................................... 137

4.2 Plasmids used in chapter four ............................................................................................ 138

4.3 Oligonucleotide primers used in chapter four ................................................................... 139

x

LIST OF FIGURES

Figure

1.1 Micrographs of C. difficile using scanning electron microscopy .......................................... 20

1.2 Micrographs of C. difficile using transmission electron microscopy ................................... 22

1.3 Pathogenicity locus (PaLoc) ................................................................................................. 24

1.4 Overview of regulators that influence toxin gene expression in C. difficile. ....................... 26

1.5 Genomic view of where toxin regulators are located in the genome relative to the pathogenicity locus. ........................................................................................................... 28

2.1 The mldABC gene cluster ..................................................................................................... 61

2.2 Sequences of Mld proteins. ................................................................................................. 63

2.3 Phenotypes of mldA::erm and mldB::erm mutants. ............................................................ 65

2.4 Complementation of mldA::erm and mldB::erm ................................................................. 67

2.5 Expression of mldA is reduced in an mldB::erm mutant ...................................................... 69

2.6 Septal localization of GFP and CFPopt fusions to division proteins ..................................... 71

2.7 Expression of CFPopt-Mld fusion proteins in wild-type and mld mutant backgrounds ...... 73

2.8 Mutants of mldA and mldB are attenuated in hamsters ..................................................... 75

2.9 Sporulation and germination ............................................................................................... 77

2.10 Toxin A production ............................................................................................................... 79

3.1. DNA sequence of mCherryOpt ........................................................................................... 104

3.2 Genetic maps of C. difficile mCherryOpt plasmids ............................................................ 106

3.3 Comparison of fluorescent proteins and autofluorescence .............................................. 108

3.4 Time course of fluorescence development ....................................................................... 110

3.5 Septal localization of mCherryOpt fusions to the division proteins MldA and ZapA ......... 112

3.6 Multiple sequence alignment of ZapA orthologs ............................................................... 114

3.7 mCherryOpt-ZapACd localizes to division sites ................................................................... 116

3.8 Use of mCherryOpt to monitor induction of the pdaV promoter by lysozyme ................. 118

3.9 Induction of PpdaV::mCherryOpt as measured by fluorescence and qRT-PCR .................... 120

xi

4.1 Schematic diagram of toxin regulation .............................................................................. 141

4.2 Bistable expression of PtcdA::mCherryOpt in C. difficile R20291 ......................................... 143

4.3 Expression of chromosomal tcdA is induced in TcdA-ON cells .......................................... 145

4.4 Alignment of tcdA promoters from different C. difficile strains ........................................ 147

4.5 Expression of PtcdA::mCherryOpt is bistable in multiple C. difficile strains ......................... 150

4.6 Effect of glucose, cysteine, and butyric acid on TcdA production ..................................... 152

4.7 PCR confirmation of C. difficile regulatory mutants .......................................................... 154

4.8 TcdR mediates bistable expression of tcdA ....................................................................... 156

4.9 Expression of PaLoc promoters in a codY null mutant ...................................................... 158

xii

LIST OF ABBREVIATIONS

ATc Anhydrotetracycline BCAAs Branched-chain amino acids C. difficile Clostridium difficile CFPopt Cyan fluorescent protein (gene is codon-optimized for C. difficile) E. coli Escherichia coli Fts Filamentous temperature sensitive GFPmut2 Green fluorescent protein variant GTP Guanosine triphosphate mCherryOpt Red fluorescent reporter (codon-optimized for C. difficile) Mld Midcell localizing division protein ORF Open reading frame PaLoc Pathogenicity island PBS Phosphate buffered saline qPCR Quantitative polymerase chain reaction qRT-PCR Quantitative real-time polymerase chain reaction RT-PCR Reverse transcriptase polymerase chain reaction Tcd Toxin of C. difficile WT Wild Type

xiii

1

CHAPTER I

INTRODUCTION

History of Clostridium difficile (Bacillus difficilis)

The story begins in the diapers of babies where in 1935 Hall and O’Toole cultured a

Gram-positive, spore-forming, motile, oxygen-sensitive bacterium (Hall and O'Toole, 1935).

They termed it Bacillus difficilis because it was rod-shaped and difficult to culture. Wondering if

B. difficilis might be pathogenic, Hall and O’Toole subcutaneously infected guinea pigs and

rabbits with a pure culture. They noticed localized edema and convulsions but no septicemia.

These convulsions were similar, but more transient, than those caused by tetanus. Sixteen of

twenty-two guinea pigs and all rabbits succumbed to infection. Hall and O’Toole were

“surprised to find the new bacillus highly pathogenic” because it was cultured from healthy

infants. To determine if B. difficile produced an exotoxin, guinea pigs were injected

subcutaneously with culture filtrate, without bacterial cells. The guinea pigs died the following

day. From this, Hall and O’Toole concluded that B. difficile produced an exotoxin, and this was

later confirmed (Snyder, 1937). In hindsight, it seems obvious that B. difficilis could be an

opportunistic pathogen to humans; however, it would be almost 40 years until it was associated

with human disease.

In the 1970s, Clostridium difficile (previously B. difficilis) was associated with diarrheal

disease and pseudomembranous colitis (described in more detail below). Pseudomembranous

colitis was initially thought to be caused by Staphylococcus aureus because this organism could

frequently be cultured from diseased patients (Khan and Hall, 1966; Smith et al., 1953). But in

1978, Bartlett et al. collected stool samples from patients with pseudomembranous colitis and

isolated a Gram-positive, strict anaerobe (Bartlett et al., 1978). The isolate was used to infect

2

hamsters, which developed a pseudomembranous colitis-like disease. Moreover, the hamsters

did not develop disease if the sample was treated with gas-gangrene antitoxin prior to infection.

These findings led Bartlett et al. to conclude that a clostridial toxin was responsible for

pseudomembranous colitis (Bartlett et al., 1978). Later in 1978, C. difficile was confirmed as an

opportunistic human pathogen (George et al., 1978; Larson et al., 1978).

Today, pseudomembranous colitis is practically synonymous with a C. difficile infection

because C. difficile is the causative agent in the vast majority of pseudomembranous colitis

cases (Farooq et al., 2015). Nevertheless, pseudomembranous colitis is occasionally linked to

other causes, including Staphylococcus aureus, Salmonella species, Clostridium ramosum,

Yersinia species, Shigella species, Campylobacter species, cytomegalovirus, Entamoeba

histolytica, inflammatory bowel disease, and certain medications (Farooq et al., 2015).

Clostridium difficile

The phylogenetic classification of C. difficile appears to be in flux. The organism belongs

to a large phylum known as the Firmicutes, which are Gram-positive bacteria with a low G+C

content. Many Firmicutes, including C. difficile, form endospores. Prominent examples of

Firmicutes include the genera Clostridia and Bacillus. Of note, Clostridia produce more protein

toxins that any other bacterial genus (Johnson, 1999). However, like many other bacteria, C.

difficile was named and classified based on phenotypic traits at a time when that approach was

standard. It has long been recognized that the taxonomy of the Clostridia based on phenotypic

traits is “clearly in need of major revision” (Collins et al., 1994).

Recently Yutin and Galperin revisited this issue using the sequences of several widely-

conserved macromolecules, namely, ribosomal proteins, the beta subunit of RNA polymerase,

DNA gyrase, and 16S rDNA (Yutin and Galperin, 2013). The analysis resulted in a thorough

3

overhaul of the Clostridia. Of note, Yutin and Galperin proposed grouping C. difficile and its

closest relatives into the family Peptostreptococcaceae and renaming C. difficile as

Peptoclostridium difficile. The new nomenclature emphasizes the relatedness to the

Peptostreptococcus species but avoids grouping a rod-shaped bacterium with

Peptostreptococcus species. However, because the name Clostridium difficile is so well-

established, it will be used throughout the remainder of this thesis.

Regarding its genetic material, several C. difficile genomes have been sequenced

(Anonymous, 2009; Darling et al., 2014; Eyre et al., 2013; Forgetta et al., 2011; Stabler et al.,

2009; van Eijk et al., 2015). The first sequenced genome was from strain CD630 (Sebaihia et al.,

2006). This genome consists of a circular chromosome of 4,290,252 bp and a native plasmid of

7,881 bp. One of the interesting findings is that a large proportion of the genome (11%) consists

of mobile genetic elements, mainly in the form of conjugative transposons.

Several typing techniques have been described for C. difficile strains, but PCR ribotyping

is the preferred method in North America. Specific ribotypes are associated with increased

sporulation titers, toxin production, and resistance mechanisms associated with antibiotics and

decontaminants (Carlson et al., 2013; Moore et al., 2013; Zidaric et al., 2012). In humans,

approximately 300 PCR ribotypes are recognized (Bauer et al., 2011). Common clinical ribotypes

include 012, 027, and 078. The ribotype 027 has emerged in the last 15 years to be the most

prominent and now accounts for 50% of isolates in North American hospitals (He et al., 2010).

The 027 genome has 50 regions of genetic difference with a total of 234 additional genes

compared with ribotype 012 (Stabler et al., 2009). Ribotype 012 is used in most laboratories

because it is more genetically malleable than other ribotypes. Ribotype 078 is another emerging

ribotype that is commonly found in bovine and porcine hosts (Keel et al., 2007). Importantly the

027 and 078 ribotypes are associated with an 18 base pair deletion within tcdC (discussed in

4

more detail below). The significance of this deletion on toxin gene expression remains unclear

(Cartman et al., 2012; Curry et al., 2007; Murray et al., 2009).

C. difficile cells are rod shaped and range from 3-17 μm in length and 0.5-1.9 μm in

width (Hafiz and Oakley, 1976; Hatheway, 1990) (Figure 1.1). As with most rod-shaped bacteria,

average length increases during log phase and decreases in stationary phase. The cell envelope

of C. difficile contains a cytoplasmic membrane, thick peptidoglycan cell wall, and a

proteinaceous S-layer. The peptidoglycan cell wall has some unusual features, most notably a

high level of N-acetylglucosamine deacetylation and mainly 3-3 cross-links (Peltier et al., 2011).

The S-layer is an essential proteinaceous layer that consists of two proteins derived from a single

polypeptide precursor, SlpA, via proteolytic cleavage (Dembek et al., 2015). To visualize the cell

envelope, our lab strain of C. difficile was grown to log phase and imaged using transmission

electron microscopy at the University of Iowa Central Microscopy Research Facility (Figure 1.2).

C. difficile is found in soil and in the intestinal flora of animals (Kim et al., 1981; McBee,

1960). Consistent with the idea that C. difficile grows primarily in mammalian hosts, the optimal

growth temperature is 37°C. C. difficile is considered motile and most strains have peritrichous

flagella (Delmee et al., 1990; Hafiz and Oakley, 1976; Tasteyre et al., 2000), with the exception

of few laboratory strains. The flagella not only provide motility but also aid in attachment in the

lower gastrointestinal tract (Tasteyre et al., 2001) and in biofilm formation (Twine et al., 2009).

C. difficile also produces a distinct smell that is attributed to three metabolic products: iso-

valeric acid, isocaproic acid, and p-cresol (Levett, 1984). Some healthcare workers note this

odor from patients with a severe C. difficile infection.

Several features of the cell envelope are important for C. difficile pathogenesis. As

noted above, flagella play a role in motility and in cell adhesion (Hafiz and Oakley, 1976;

Tasteyre et al., 2001). While most C. difficile produce flagella, some lab strains do not, so

5

studying pathogenesis in these strains can be misleading (Purdy et al., 2002; Tasteyre et al.,

2001). Some strains of C. difficile are thought to produce a capsule that assists with evasion

from the immune system (Davies and Borriello, 1990). The extensive deacetylation of the

peptidoglycan backbone contributes to lysozyme resistance (Ho et al., 2014), while the

preference for 3-3 crosslinks over 3-4 crosslinks probably contributes to resistance against β-

lactam antibiotics (Bera et al., 2005). Lastly, the S-layer is thought to aid in attachment to host

cells and interact with the immune system (Spigaglia et al., 2013).

C. difficile is considered a strict anaerobe (Edwards et al., 2013; Winn and Koneman,

2006). Survival of vegetative cells is limited to anaerobic environments, including the

gastrointestinal tract. Fortunately for C. difficile, it can differentiate into a metabolically-inert

and aerotolerant spore. The ability to form spores is vital for transmission (Lawley et al., 2009).

As a spore, C. difficile is resistant to some disinfectants, cleaners, and alcohol-based sanitizers

(Lawley et al., 2010; Wullt et al., 2003), which contributes to the high rates of nosocomial

infection (Dubberke et al., 2007; Riggs et al., 2007). These spores also can allow C. difficile to

remain dormant and avoid competing with normal flora in the lower gastrointestinal tract

(Borriello, 1990; Camorlingaponce et al., 1985). Once the normal flora is disrupted, the spores

can germinate and the vegetative cells can prosper. In the laboratory sodium taurocholate, a

component of bile, is used to germinate spores (Wilson et al., 1982). C. difficile is sensitive to

some secondary bile metabolites and recent data suggest that these may be how the

microbiome controls C. difficile infection (Greathouse et al., 2015).

Epidemiology

The colonization rate of C. difficile varies depending on health status and age

(Cooperstock et al., 1983; Tullus et al., 1989). The numbers vary from 4-15% in healthy adults to

6

>70% in infants (Bartlett, 1994; Kato et al., 2001). Colonization is less concerning in infants

because disease manifestation is rare. It is hypothesized that infants lack the receptors needed

for toxin entry and therefore colonization does not result in disease (Schaffer et al., 1987). Even

in adults who have the receptor, most people who carry C. difficile remain asymptomatic.

C. difficile is the most common cause of nosocomial diarrhea in developed countries

(Bacci et al., 2009; Kuipers and Surawicz, 2008). In addition, nearly one in five cases of antibiotic

associated diarrhea result from C. difficile infection (Bartlett and Gerding, 2008). In the United

States, C. difficile is linked to >14,000 deaths and >250,000 infections each year (CDC, 2013).

These numbers are slightly down from the peak in 2011 when almost 500,000 infections were

reported. Thus, C. difficile is the second most frequently isolated enteric pathogen, behind

Campylobacter jejuni (Indra et al., 2009).

The Centers for Disease Control and Prevention have identified C. difficile as an “urgent”

threat to public health (CDC, 2013). Only two other bacteria were categorized in this highest

threat level, carbapenem-resistant Enterobacteriaceae and drug-resistant Neisseria

gonorrhoeae. It is vital to study and monitor these pathogens as they are a serious threat to

public health. Probably the most terrifying statistic regarding C. difficile infections is the high

relapse rate. Approximately 20% of symptomatic patients will experience a relapse (Louie et al.,

2011; Lowy et al., 2010). Relapse is defined operationally as a recurrence of infection within 28

days since completion of the previous antibiotic therapy, whereas an infection that appears

more than 28 days later is regarded as a re-infection. The probability of relapse or re-infection

doubles for each subsequent infection (Comely et al., 2012), and the prognosis for patients with

recurring C. difficile infections is poor.

Risk factors for a C. difficile infection include antibiotic therapies, treatment with

proton-pump inhibitors, age, and hospital exposure (Barbut and Petit, 2001; Eyre et al., 2012).

7

The single biggest risk factor is previous antibiotic therapy, especially third generation

cephalosporins, fluoroquinolones, and clindamycin (Buffie et al., 2012; Chang et al., 1978; Chen

et al., 2008; Delalla et al., 1989; Gerding, 2004). These antibiotics disrupt the normal flora and

thus create a niche for the opportunistic C. difficile to flourish. Why proton-pump inhibitors are

major risk factors is unclear (Daniell, 2014; Tleyjeh et al., 2012; Yearsley et al., 2006). One

hypothesis is that a more neutral pH in the gastrointestinal tract allows survival of vegetative C.

difficile cells and alters the normal flora. Lastly, elderly persons and immunocompromised

individuals are more susceptible to C. difficile infections (Jagai and Naumova, 2009; Nakamura et

al., 1981; Tal et al., 2002; Treloar and Kalra, 1987). This is likely due to a combination of factors:

high prevalence of antibiotic usage, waning immunity, decreased biodiversity of the intestinal

microbiome, and time spent in hospitals and nursing homes.

Clinical aspects of Clostridium difficile

The most common symptom of a C. difficile infection is diarrhea. Additional symptoms

can include abdominal pain, fever, nausea, and dehydration (Chang et al., 2007; Kyne et al.,

1999). If the patient experiences significant fluid loss, rehydration therapy is administered.

Antibiotic therapy is a frequent treatment strategy as well. The most common antibiotics for C.

difficile infections include metronidazole for first time infections and vancomycin for recurrent

infections (Cohen et al., 2010; McFarland et al., 2002). Certain antibiotics (third generation

cephalosporins, fluoroquinolones, and clindamycin) are not only ineffective but are considered

risk factors for C. difficile infections (Buffie et al., 2012; Chang et al., 1978; Chen et al., 2008;

Delalla et al., 1989; Gerding, 2004).

Pseudomembranous colitis is an acute inflammatory disease of the colon where plaques

form on the intestinal walls. These plaques consist of leukocytes and appear yellow or white.

8

Although pseudomembranous colitis is often treatable, it can be life-threatening (Ruiz et al.,

2015). Treatments include antibiotic therapy and/or surgical intervention. Pseudomembranous

colitis is highly associated with antibiotic use, with symptoms of diarrhea beginning any time

from the first days of antibiotic treatment to several weeks after the antibiotic treatment is over

(Han et al., 2009).

C. difficile infections can progress to life-threatening conditions like toxic megacolon and

colonic perforation (Velanovich et al., 1992). Toxic megacolon is colon inflammation and

dilation, which may prevent peristalsis or food movement. Colonic perforation is a

gastrointestinal breach and is life-threatening (Rausch and Kaibie, 2014). For both conditions,

sepsis and shock often occur and usually require a colectomy (Panis et al., 1992). Even with

surgery, the mortality rate in patients with toxic megacolon is 35% (Mitas et al., 2012).

C. difficile has on occasion been found beyond the gastrointestinal tract (Choi et al.,

2013; Cid et al., 1998; Feldman et al., 1995; Hemminger et al., 2011; Kaufman et al., 2013; Libby

and Bearman, 2009; Quera et al., 2014). However, this is quite rare and the patients are often

severely immune comprised or near death. When the patient is near death, the colon may have

already died. This causes collapse of the colon walls, allowing microbes to invade the

surrounding tissue and bloodstream.

Recently there has been a lot of interest using fecal replacement therapy to treat

recalcitrant cases of recurring C. difficile infection (Boyle et al., 2015; Kelly et al., 2014; Patel et

al., 2013; Stollman and Surawicz, 2012). The basis for this approach is that restoration of the

normal flora provides an instant and effective microbial competition capable of displacing C.

difficile. In practical terms, fecal replacement therapy involves administering a homogenized

stool sample from a matched donor such as a household partner or relative directly into the

gastrointestinal tract via colonoscopy. While not yet FDA approved, preliminary data suggest

9

this is an effective method of treatment and noticeable improvement is observed in less than 48

hours.

One terrifying truth about fecal replacement therapy is that little is known about the

microbial composition of the donor material, creating the worrisome possibility of introducing

new pathogens into an already deathly ill patient. To prevent this from happening, some

institutions have begun screening the donor feces with a standard stool pathogen panel.

Unfortunately, the out-of-pocket expense is several thousand dollars and insurance rarely

covers this testing since the donor is healthy. One hopes that in the future the stool may be

screened similar to blood prior to transfusions to ensure the patients receive harmless microbes

and not pathogens. Alternatively, there is work towards developing a defined cocktail of

characterized microbes that can be used in lieu of an undefined fecal transplant (Rohlke and

Stollman, 2012).

There is at least one report of a fecal transplant causing harm to the recipient (Weil and

Hohmann, 2015). In this case, a 32-year-old woman with recurrent and life-threatening C.

difficile infections received a fecal transplant from her 16-year-old daughter. At the time, the

daughter exhibited no health problems, had normal weight, and a screen of her stool did not

reveal the presence of any pathogens. Curiously, 16 months after the transplant, both the

mother and daughter gained and maintained significant weight, even after a diet and exercise

program. The cause of the weight gain is not clear, but this case has been interpreted to mean

that the daughter already had a microbiome that predisposed her to obesity, which was

transferred to the mother. Whatever the true basis of this therapeutic outcome, the case is a

stark reminder that fecal replacement therapy is not without potential risks.

Clinical diagnosis

10

C. difficile can be diagnosed using an immunoassay test, polymerase chain reaction

(PCR), colonoscopy, and advanced imaging like a computerized tomography scan (Planche and

Wilcox, 2015). The clinical microbiology laboratory at the University of Iowa Hospitals and

Clinics utilizes a new diagnostic algorithm with three components. A sample is first tested for

the presence of toxin antigen using an immunoassay test. If positive, the patient is considered

to have a C. difficile infection. If negative, the sample is tested using a glutamate

dehydrogenase test. A positive test suggests C. difficile (toxigenic or non-toxigenic) is present,

but is not definitive because glutamate dehydrogenase is not unique to C. difficile. Patients who

test glutamate dehydrogenase positive and toxin negative are subsequently tested using a PCR

for toxin genes. Collectively, these assays discriminate between toxigenic and non-toxigenic C.

difficile strains and provide physicians with reliable test results without excessive costs owing to

performing too many diagnostic procedures.

Toxins of C. difficile

C. difficile pathogenesis is mediated primarily by Toxin A (TcdA) and Toxin B (TcdB).

TcdA and TcdB are thought to be the main contributors to C. difficile virulence based on several

lines of evidence. Firstly, toxin genes are required for virulence in animal models (Kuehne et al.,

2010; Kuehne et al., 2014; Lyras et al., 2009). Secondly, antibody-mediated therapies that target

the toxins improve prognosis in animal models (Babcock et al., 2006; Steele et al., 2013).

Thirdly, administration of purified toxins recapitulates disease in animal models (Lyerly et al.,

1985; Triadafilopoulos et al., 1987). Lastly, 99.5% of clinical C. difficile isolates encode at least

one toxin (Eckert et al., 2015). In clinical isolates, TcdB is almost always present (Voth and

Ballard, 2005); however, TcdA can be absent in these strains. Interestingly, a clinical isolate that

is TcdA-positive and TcdB-negative has not been reported.

11

TcdA and TcdB are quite similar and likely arose from a duplication event (von Eichel-

Streiber et al., 1992). Both toxins contain four domains: N-terminal glucosyltransferase domain,

an autocatalytic cysteine proteinase domain, a central translocation domain with a hydrophobic

region, and a C-terminal receptor binding domain (Jank et al., 2007a, b; Voth and Ballard, 2005).

The total size of TcdA and TcdB is 308 and 269 kDa, respectively (Pothoulakis and Lamont, 2001;

Savidge et al., 2003), making these proteins among the largest known bacterial toxins.

TcdA and TcdB are internalized using a dynamin-dependent process that is mainly

governed by clathrin (Papatheodorou et al., 2010). However, the specific receptor that

mediates entry appears to differ for each toxin. The receptor for TcdA is thought to be the

disaccharide GalΒ1-4GlcNac (Gerhard et al., 2013; Greco et al., 2006; Na et al., 2008; Wilkins and

Tucker, 1989). On the other hand, TcdB utilizes chondroitin sulfate proteoglycan 4 (Yuan et al.,

2015). Following receptor-mediated endocytosis, toxins traffic to a low pH endosome where

conformational changes (Qa'Dan et al., 2000) activate the intrinsic cysteine protease activity

(Genisyuerek et al., 2011) to release the glucosyltransferase domain. The toxin works by

transferring glucose from UDP-glucose to several host proteins, especially the Rho family of

GTPases (Just et al., 1995a). The Rho GTPases play important roles in regulating dynamics of the

actin cytoskeleton and in several signal transduction pathways. Covalent modification of the

Rho GTPases inactivates these proteins, leading to cytoskeletal collapse, cell rounding, loss of

tight junctions, apoptosis, and inflammation (Just et al., 1995b; Just et al., 1995c; Poxton et al.,

2001). This localized distress results in an influx of neutrophils, resulting in the hallmark

symptom pseudomembranous colitis (Kelly et al., 1994). In addition to the glucosyltransferase

activity, TcdB has been shown to kill epithelial cells by a glucosyltransferase-independent

mechanism (Chumbler et al., 2012; Wohlan et al., 2014).

12

Some strains of C. difficile produce a third known as the binary toxin. This is an actin-

specific ADP-ribosyltransferase that causes depolymerization of the actin cytoskeleton (Perelle

et al., 1997; Popoff et al., 1988). As the name suggests, binary toxin comprises two subunits, an

enzymatic component and a binding component that uses the host lipolysis-stimulated

lipoprotein receptor (Papatheodorou et al., 2011). Although binary toxin is cytopathic to cells in

culture, it is thought play a minor role in causing disease in humans. Binary toxin can be

detected in ~20% of clinical C. difficile isolates (Barbut et al., 2007; Bauer et al., 2011; Geric et

al., 2006; Geric et al., 2003; Stubbs et al., 2000) and has been associated with exacerbated

disease (Bacci et al., 2011; Barbut et al., 2005). Interestingly, there is a report of clinical C.

difficile isolates producing only binary toxin (Geric et al., 2003).

Pathogenicity Locus (PaLoc)

The genes encoding Toxin A (tcdA) and Toxin B (tcdB) are located on a 19.6 kb

pathogenicity locus (PaLoc) (Braun et al., 1996), along with three accessory genes: tcdE, tcdR,

and tcdC (Figure 1.3). Of the three accessory genes, only the function of tcdR is undisputed.

TcdR is an alternative sigma factor that directs RNA polymerase to the promoters for tcdA, tcdB,

and tcdR itself (Mani and Dupuy, 2001; Mani et al., 2002). Thus, TcdR is required for toxin

production (Mani and Dupuy, 2001). As will be discussed below, TcdR is thought to be a major

focal point for regulation of toxin production.

The function of tcdC is a matter of contention. Some studies indicate TcdC is a negative

regulator of toxin production, perhaps functioning as an anti-sigma for TcdR. This notion is

supported by reports of elevated toxin production in clinical isolates with deletions or frame-

shift mutations predicted to inactivate tcdC (Curry et al., 2007; Warny et al., 2005). Moreover,

in vitro TcdC interferes with the function of RNA polymerase containing sigma TcdR but not with

13

the function of RNA polymerase containing sigma A (Matamouros et al., 2007). However,

according to other reports, there is no clear correlation between a tcdC genotype and toxin

production, and insertional inactivation of tcdC does not increase toxin production (Cartman et

al., 2012; Rosenbusch et al., 2012). At present there is no satisfactory explanation for these

conflicting results.

The function of the final PaLoc gene, tcdE, is also unsettled. TcdE is 139 amino acids

long and has three strongly-predicted transmembrane helices. It has been suggested that TcdE

is similar to pore-forming “holin” proteins and thus facilitates toxin secretion (Tan et al., 2001).

The question of how Toxin A and Toxin B are released to the environment is an interesting one,

because neither toxin has an obvious signal sequence for export. Unfortunately, the two

published studies that tested the role of TcdE in toxin secretion came to diametrically opposed

conclusions (Govind and Dupuy, 2012; Olling et al., 2012).

The G+C content of the PaLoc is lower than that of the chromosome in general (26% vs.

29%), suggesting it was acquired by horizontal gene transfer. C. difficile strains that lack the

PaLoc are “non-toxigenic,” though they may produce binary toxin, which is not encoded on the

PaLoc. In these non-toxigenic strains, a 115 bp sequence occurs at the site of the PaLoc.

Although it has been suggested that the PaLoc has some similarities to mobile genetic elements,

it is always found at the same insertion site (when it is present at all), and the ends of the PaLoc

do not display sequences characteristic of transposons. Strain-to-strain differences in the DNA

sequence of the PaLoc have been used to establish 32 toxinotypes of C. difficile (Zidaric et al.,

2008).

Regulation of toxin expression by metabolic inputs

14

Because of the central role that Toxins A and B play in C. difficile pathogenesis, there has

been a lot effort invested in understanding how toxin production is regulated. For a recent

review, see Bouillaut et al. (2014). Transcription of the toxin genes is repressed during

exponential phase and induced upon entry into stationary phase, suggesting toxin production is

a response to limitation for nutrients and/or energy (Dupuy and Sonenshein, 1998; Hundsberger

et al., 1997). Consistent with this notion, supplementing the growth media with glucose impairs

toxin production during entry into stationary phase (Antunes et al., 2012; Dupuy and

Sonenshein, 1998). Other inhibitors of toxin production include cysteine, branched chain amino

acids, proline, and butanol (Karlsson et al., 2008; Karlsson et al., 2003; Karlsson et al., 2000).

The optimal temperature for toxin production is 37°C (Karlsson et al., 2003). Collectively, these

studies indicate toxin production has evolved as a mechanism for procuring nutrients from a

mammalian host.

The master regulator for tcdA and tcdB expression is the alternative sigma encoded by

tcdR (Hundsberger et al., 1997). Expression of tcdR parallels expression of the toxin genes and

thus responds to the same metabolic inputs that modulate toxin production (Antunes et al.,

2012; Karlsson et al., 2008; Karlsson et al., 2003; Karlsson et al., 2000). Besides activating

expression of tcdA and tcdB, TcdR activates its own expression, which creates a positive

feedback loop that can greatly amplify an inducing signal (Moncrief et al., 1997). As noted

above, some reports indicate TcdC is a negative regulator of toxin production that works by

inhibiting the activity of TcdR (Curry et al., 2007; Dupuy et al., 2008; Matamouros et al., 2007),

but other studies have challenged this proposal (Carter et al., 2011b; Cartman et al., 2012;

Murray et al., 2009).

A multitude of global regulators have been implicated in coupling expression of tcdA

and tcdB to the metabolic status of the cell. These include CodY, CcpA, PrdD, Rex, SigD, Spo0A,

15

and SigH. These genes for these factors are not found on the PaLoc but instead are distributed

throughout the chromosome (Figure 1.4).

CodY, a global regulator of transcription found in a wide variety of Firmicutes, has been

studied most extensively in Bacillus subtilis, where it represses expression of hundreds of genes

in nutrient-rich conditions (Belitsky and Sonenshein, 2013). The DNA-binding activity of CodY is

controlled by two classes of metabolites that bind directly to CodY and act synergistically: GTP

and branched-chain amino acids (isoleucine, leucine, and valine). GTP is an indicator of the

energy status of the cells while BCAAs convey information about the nutritional status of the

cell. During entry into stationary phase, decreases in the cellular pools of GTP and BCAAs trigger

release of CodY from its DNA-bind sites, which activates expression of hundreds of genes

(Ratnayake-Lecamwasam et al., 2001). Many of these genes have clear connections to

metabolism, e.g., they encode proteins that help the cell assimilate nutrients, synthesize

building blocks such as amino acids, or are involved in sporulation.

In C. difficile, transcriptional profiling indicates CodY represses expression of 146 genes

and activates expression of 19 genes (Dineen et al., 2010). Although most of the CodY-regulated

genes fall into the same functional classes as observed in B. subtilis, one notable exception is

that in C. difficile CodY represses expression of all of the genes on the PaLoc (Dineen et al.,

2007). CodY has previously been implicated in regulating expression of toxins in other

Firmicutes (Majerczyk et al., 2008; van Schaik et al., 2009). Biochemical studies revealed that C.

difficile CodY binds with high affinity to the promoter region of the sigma tcdR and with lower

affinity to the promoter regions of tcdA and tcdB. Thus, CodY probably represses toxin

production primarily by repressing expression of tcdR during exponential growth, when levels of

GTP and BCAAs are relatively high. CodY is also likely responsible for mediating the repression

16

of toxin production observed when the growth medium is spiked with BCAAs (Karlsson et al.,

2008).

Catabolite control protein A (CcpA) is a DNA-binding protein that acts as a global

regulator of carbon metabolism in Firmicutes (Fujita, 2009; Johnson et al., 2009; Mahr et al.,

2002). Studies done primarily with CcpA from B. subtilis indicate that the DNA binding activity of

CcpA is allosterically regulated by several ligands, the most important being a form of the Hpr

protein that is phosphorylated on Serine 46 (Hpr-Ser46-P) (Schumacher et al., 2004).

Phosphorylation of Hpr on Ser46 is catalyzed by Hpr kinase/phosphorylase, whose activity is

stimulated by glycolytic intermediates such as fructose-1,6-bisphosphate (Deutscher and Saier,

1983). Thus, Hpr-Ser46-P accumulates under glucose replete conditions when intermediates of

glycolysis are relatively abundant. [The regulatory function of Hpr is distinct from its perhaps

better-known role as part of the phosphotransferase system for sugar uptake; when functioning

in that capacity Hpr is phosphorylated on His15 (Horstmann et al., 2007)].

In C. difficile, CcpA regulates expression of 10% of the genome (Antunes et al., 2012).

About 140 of these genes are thought to be regulated directly based on the identification of

CcpA binding sites at these genes (Antunes et al., 2012). As expected, many CcpA-regulated

genes are involved in fermentation, sugar transport, and amino acid metabolism, but several

genes on the PaLoc are also subject to repression by CcpA: tcdA, tcdB, tcdR, and tcdC (Antunes

et al., 2012; Antunes et al., 2011a). CcpA binds directly to the promoter regions for each of

these genes (Antunes et al., 2011a). In a ccpA mutant, toxin production is no longer repressed

by exogenous glucose, indicating that CcpA mediates glucose-repression of toxin synthesis.

Curiously, toxin production is also lower in the absence of glucose in a ccpA mutant, an

observation that is not easily accounted for based on the known functions of CcpA.

Interestingly, in a departure from the B. subtilis paradigm, in C. difficile fructose-1,6-

17

bisphosphate rather than Hpr-Ser46-P is the ligand that binds to CcpA to potentiate DNA-

binding (Antunes et al., 2012). Moreover, CcpA from B. subtilis and C. difficile have a different

consensus binding site (Antunes et al., 2012).

SigD is an alternative sigma factor that is found in some Gram-negative and Gram-

positive bacteria (Colland et al., 2001; Helmann and Chamberlin, 1987; Marquez et al., 1990).

SigD has been shown to positively regulate the expression of flagellar genes in a hierarchical

manner in Salmonella enterica, Pseudomonas aeruginosa, and B. subtilis (Chilcott and Hughes,

2000; Mirel and Chamberlin, 1989; Starnbach and Lory, 1992). In C. difficile, SigD responds to

intracellular concentrations of the second messenger cyclic diguanylate and then alters late-

stage flagellar genes (McKee et al., 2013; Purcell et al., 2012). Moreover, SigD was identified as

a key regulator of virulence genes in C. difficile (Aubry et al., 2012; McKee et al., 2013). SigD

increases toxin gene expression by directly increasing the expression of tcdR, the master

regulator of toxin (El Meouche et al., 2013).

PrdR and Rex are thought to link amino acid fermentation to toxin production. PrdR

activates expression of proline reductase gene cluster (Bouillaut et al., 2013; Jackson et al.,

2006). Proline represses toxin production and is a major electron acceptor during amino acid

fermentation. It is thought that in the absence of proline alternative pathways, regeneration of

NAD+ from NADH must be pressed into service, but these are relatively inefficient, so the

NADH/NAD+ ratio increases. Rex is a widely conserved regulator of anaerobic metabolism that

can bind both NAD+ and NADH (Sickmier et al., 2005). But there is an important difference—

only the Rex:NAD+ complex binds DNA and represses expression of target genes. By sensing the

NADH/NAD+ ratio, Rex monitors not only the redox balance of the cell but also whether

fermentation (and thus generation of ATP) is operating efficiently. In the absence of proline or

other suitable electron acceptors, NADH levels rise, triggering derepression of toxin production,

18

and other pathways under Rex-control. Because binding sites for PrdR or Rex have yet to be

identified in the PaLoc, it seems likely that PrdR and Rex regulate toxin production indirectly.

Spo0A is a response regulator, best known for activating the sporulation cascade in B.

subtilis (Olmedo et al., 1990). Spo0A has also been shown to regulate competence,

transformation, DNA replication, biofilm formation, virulence factors, stress responses, and

solvent production (Castilla-Llorente et al., 2006; Chen et al., 2010; Hahn et al., 1995; Hamon

and Lazazzera, 2001; Ravagnani et al., 2000; Saile and Koehler, 2002). In C. difficile, Spo0A has

been shown to regulate sporulation, metabolism, and biofilm formation (Dawson et al., 2012;

Deakin et al., 2012; Pettit et al., 2014). Spo0A has also been shown to regulate toxin gene

expression (Mackin et al., 2013; Rosenbusch et al., 2012); however, this may be strain specific

since the effect on toxin gene expression is observed in ribotypes 027 but not ribotypes 078 or

012 (Mackin et al., 2013). Spo0A likely influences toxin gene expression indirectly because no

binding sites have been found in the PaLoc.

SigH (Spo0H) is an alternative sigma factor found in most Firmicutes. The SigH regulon

in B. subtilis includes genes involved in sporulation, biofilm formation, cell wall metabolism,

proteolysis, and many other factors involved in adapting to nutrient depletion (Asai et al., 1995;

Britton et al., 2002; Hamon et al., 2004). SigH has also been shown to regulate toxin synthesis in

Bacillus anthracis (Hadjifrangiskou et al., 2007). Transcription of sigH is controlled directly by

the transcriptional repressor AbrB and indirectly by the phosphorylated form of Spo0A

(Spo0A∼P), which represses abrB (Perego et al., 1988; Weir et al., 1991). SigH also regulates its

own expression. SigH activity has been shown to respond to carbon source (Dixon et al., 2001)

and pH (Cosby and Zuber, 1997). In C. difficile, SigH influences sporulation, metabolism, and

toxin production (Karlsson et al., 2008; Saujet et al., 2011). In a sigH null mutant, the expression

of tcdR, tcdA, and tcdB increased. It is unlikely that SigH is directly regulating expression of

19

these genes because the PaLoc lacks any obvious SigH binding sites (Rosenbusch et al., 2012;

Saujet et al., 2011).

In summary, toxin production appears to be a response to nutrient limitation. At least

in the laboratory, the timing and extent of toxin production can be manipulated by altering the

growth medium. Moreover, most of the regulatory proteins that appear to control toxin

production are also involved in adaptation to various nutritional stresses.

Overview of chapters in this thesis

Chapter 2 describes the characterization of three proteins that are essentially unique to

C. difficile. Before our studies, these proteins had no known function. We found that all three

proteins are midcell localizing division proteins and therefore were named MldA, MldB, and

MldC. The Mld proteins play a role in cell division, cell shape, and pathogenesis. The work in

this chapter has been published (Ransom et al., 2014).

Chapter 3 describes a new oxygen-dependent red fluorescent protein named

mCherryOpt, which was codon-optimized for C. difficile. This reporter can be used to study

protein localization at a subcellular level and study gene expression in an individual cell. The

findings described in chapter 3 have been published (Ransom et al., 2015).

In chapter 4, we report that toxin gene expression C. difficile is bistable. In addition, we

found this bistability is dependent upon the master regulator of toxin production, TcdR. With

this new insight and the mCherryOpt reporter, we revisit toxin regulation to confirm existing

paradigms and propose a revised model for C. difficile toxin regulation.

Finally, chapter 5 describes future directions for continued studies of this work.

20

21

Figure 1.1. Micrograph of C. difficile using scanning electron microscopy. Cells were grown to

log phase in broth media and imaged. Furrows indicate sites of cell division.

22

23

Figure 1.2. Micrograph of C. difficile using transmission electron microscopy. Cells were grown

to log phase and imaged. (A) Cell pole. (B) Division site.

24

25

Figure 1.3. Pathogenicity locus (PaLoc). This region encodes for the two main toxins (TcdA and

TcdB) as well as the master regulator of toxin TcdR. The PaLoc also encodes for TcdE and TcdC.

TcdE may be involved in toxin release, and TcdC may act as an anti-sigma factor against TcdR.

The numbers indicate the distance between the respective genes.

26

Toxin gene expression

Spo0A SigH

SigD

PrdR Rex

CodY CcpA

27

Figure 1.4. Overview of regulators that influence toxin gene expression in C. difficile.

28

29

Figure 1.5. Genomic view of where toxin regulators are located in the genome relative to the

pathogenicity locus. The genomic locus of the mldABC genes is also indicated. Color indicates

the GC percentage in 10,725 base pair increments. C. difficile genomes include a native plasmid

as well but it is not shown.

30

CHAPTER II

Identification and characterization of a gene cluster required for proper rod shape, cell division, and pathogenesis in Clostridium difficile

Introduction

Clostridium difficile is a strictly anaerobic, Gram positive, spore-forming bacterium that

has become the leading cause of hospital-acquired diarrhea in developed countries. The annual

impact of C. difficile infections in the United States has been estimated at 14,000 deaths and

over $1 billion in excess medical costs (CDC, 2013). Both the severity and the frequency of C.

difficile infections are increasing (Kelly and LaMont, 2008), and a recent report on the impact of

antibiotic resistance classified the organism as an “urgent threat,” the highest threat level (CDC,

2013).

C. difficile infections typically occur in people who have been treated with antibiotics

that disrupt the flora of the gastrointestinal tract (Owens et al., 2008; van Nood et al., 2013).

Although C. difficile is resistant to many antibiotics, the infection usually resolves upon

treatment with metronidazole or oral vancomycin (Nelson et al., 2011). Unfortunately, disease

recurs in ~20% of patients, and for this cohort the prognosis is poor (Fekety et al., 1997; O'Horo

et al., 2013; van Nood et al., 2013). The high rate of recurrence has been attributed to

germination of C. difficile spores after antibiotic therapy is ended, but before restoration of the

normal flora (Carroll and Bartlett, 2011; Kelly and LaMont, 2008). For this reason there is

interest in developing antibiotics that target C. difficile selectively and in treatments like fecal

transplants which work by restoring a healthy microbiome (Chilton et al., 2013; Goldstein et al.,

2005; O'Horo et al., 2013; van Nood et al., 2013).

Here we describe a three-gene operon found in C. difficile that is important for

morphogenesis, cell division, and pathogenesis. We named the genes in this operon mldA,

31

mldB, and mldC, since they encode midcell localizing division proteins. Because these genes are

restricted to C. difficile and a few of its closest relatives, drugs that inhibit the Mld proteins

might target C. difficile without disrupting the intestinal microbiome. We also describe a

method for using green fluorescent protein (GFP) and cyan fluorescent protein (CFP) to study

protein localization in strict anaerobes. Because the various color variants of GFP all require O2

for chromophore development (Tsien, 1998), their use has been largely restricted to aerobic

bacteria or to anaerobes that tolerate transient (e.g., 20 min.) exposure to air (Hartman et al.,

2011). The extension of O2-dependent fluorescent proteins to strict anaerobes should facilitate

studies of protein localization, gene expression and high-throughput screens for antibiotics in

this very important class of bacteria.

Materials and Methods

Strains, media, and growth conditions

Bacterial strains used in this study are listed in Table 2.1. All C. difficile strains are

derived from the erythromycin sensitive isolate JIR8094, which is in turn derived from the

sequenced strain CD630 (O'Connor et al., 2006; Sebaihia et al., 2006). C. difficile was routinely

grown in Tryptone Yeast (TY) or Brain Heart Infusion (BHI) media, supplemented as needed with

thiamphenicol at 10 μg/ml, erythromycin at 5 μg/ml, kanamycin at 50 μg/ml, or cefoxitin at 16

μg/ml. TY consisted of 0.4% tryptone, 0.5% yeast extract, 0.1% L-cysteine and 0.5% NaCl (where

indicated). BHI consisted of 3.7% brain heart infusion medium (Gibco) supplemented with 0.5%

yeast extract, 0.4% glucose, and 0.1% L-cysteine. For solid media, agar was added at 2% final

concentration. C. difficile spores were germinated on cycloserine cefoxitin fructose agar (CCFA)

plates containing 1.5% agar, 4% protease peptone, 0.5% sodium phosphate dibasic, 0.1%

monopotassium phosphate, 0.2% NaCl, neutral red, 0.006% magnesium sulfate, 0.6% fructose,

32

0.1% L-cysteine, 16 μg/ml cefoxitin, 125 μg/ml cycloserine, and 0.1% taurocholate (Wilson et al.,

1982). C. difficile strains were maintained at 37°C in an anaerobic chamber (Coy Laboratory

products) in an atmosphere of 10% H2, 5% CO2, and 85% N2.

Escherichia coli strains were grown in LB medium at 37°C with ampicillin at 200 μg/ml or

chloramphenicol at 20 μg/ml as needed. LB contained 10% tryptone, 5% yeast extract, 1% NaCl,

and, for plates, 1.5% agar.

Plasmid and strain construction

The oligonucleotide primers used in this work are listed in Table 2.2 and were

synthesized by Integrated DNA Technologies (Coralville, IA). All plasmids were verified by DNA

sequencing and are listed in Table 2.3.

C. difficile null mutants of mldA and mldB were constructed using modified TargeTron

procedures (Sigma-Aldrich) to insert a group II intron conferring Erm resistance (Heap et al.,

2010). Primers for retargeting the group II intron were designed using algorithms generously

provided by Rob Britton (Michigan State University) or ClosTron (Heap et al., 2010). First, an E.

coli-C. difficile shuttle vector designated pTHE1037 was constructed by digesting pMC123

(McBride and Sonenshein, 2011) with BspHI and self-ligating, which results in loss of bla. To

retarget the intron to insert after nucleotide 248 of mldA, the intron template was amplified by

PCR as outlined in the TargeTron user manual (Sigma-Aldrich) using the EBS universal primer

CDE914 in combination with primers P1447, P1448, and P1449. The resulting PCR product was

digested with HindIII and BsrG1, then ligated into pCE240 (Ho and Ellermeier, 2011) that had

been digested with the same enzymes, resulting in pDSW1215. Then pDSW1215 was digested

with SphI and SfoI to obtain a TargeTron fragment, which was cloned into pTHE1037 that had

been digested with EcoRI, made blunt-ended using Klenow fragment, and then digested with

33

SphI. The resulting plasmid was designated pRAN101. A plasmid that targets the group II intron

to nucleotide 153 of mldB was constructed similarly except: the primer set was CDE914, RP114,

RP115, and RP116. The PCR product was cloned onto HindIII-BsrG1-digested pBL100 (Bouillaut

et al., 2013), resulting in pRAN243.

Plasmids pRAN101 (mldA248) and pRAN243 (mldB153) were transformed into the E. coli

conjugation donor strain HB101/pRK24 (Trieu-Cuot et al., 1987). Then pRAN101 and pRAN243

were transferred to C. difficile JIR8094 via conjugation and selection for Erm-resistance as

described (Heap et al., 2010; Ho and Ellermeier, 2011). The insertion of introns into mldA and

mldB were confirmed by PCR using primer pairs RP1/RP2 (mldA248) or RP28/RP29 (mldB153).

Finally, loss of the TargeTron plasmid was confirmed by thiamphenicol sensitivity.

For complementation studies we constructed a set of plasmids that carry various

combinations of mldABC. The plasmids are all derivatives of pRPF185, which has a tetracycline-

inducible promoter (Fagan and Fairweather, 2011). Genes were amplified using the following

primer sets: mldABC (RP164 and RP165), mldAB (RP164 and RP187), mldA (RP164 and RP186),

mldBC (RP184 and RP165), mldB (RP184 and RP187), and mldC (RP185 and RP165). The

resulting PCR products were digested with SacI and BamHI then ligated into pRPF185 digested

with the same enzymes. The resulting plasmids were designated pRAN414 (mldABC), pRAN415

(mldAB), pRAN416 (mldA), pRAN417 (mldBC), pRAN418 (mldB) and pRAN419 (mldC). They were

introduced into C. difficile strains by conjugation from HB101/pRK24, selecting for

thiamphenicol resistance.

To study the localization of Mld proteins, we constructed gene fusions to a low GC

codon-optimized variant of cyan fluorescent protein, CFPopt (Sastalla et al., 2009). A basic

CFPopt expression vector was constructed by using PCR to amplify cfpopt from template pSW4-

CFPopt (Sastalla et al., 2009) with primers RP160 and RP161. The resulting PCR product was

34

digested with SacI and BamHI, and ligated into pRPF185 that had been cut with the same

enzymes. The resulting plasmid, pRAN334, expresses CFPopt followed by a stop codon under

control of a tetracycline-inducible promoter. A plasmid designated pRAN357, in which the

coding sequence of CFPopt is followed by a multicloning site for making fusions of CFPopt to the

N-terminus of target proteins, was constructed similarly using primers RP161 and RP171.

CFPopt–Mld fusions were constructed by PCR amplifying the mld genes using the following

primer pairs: mldA (RP178-RP179), mldB (RP196-RP197), or mldC (RP198-RP199). The resulting

PCR products were digested with SphI and AscI and cloned onto pRAN357 digested with the

same enzymes resulting in CFPopt-MldA (pRAN410), CFPopt-MldB (pRAN460), and CFPopt-MldC

(pRAN461). These plasmids were introduced into C. difficile by conjugation from HB101/pRK24,

selecting for thiamphenicol resistance.

Intergenic RT-PCR and qRT-PCR

RNA was harvested from logarithmically growing cells using the RNeasy system

(Qiagen). The Access RT-PCR system (Promega) was used for reverse transcriptase PCR (RT-

PCR). Quantitative real-time PCR (qRT-PCR) experiments were performed as previously

described (18) using Sybr green master mix (Applied Biosystems) and the following gene-specific

primer pairs: mldA (RP34-RP35), mldB (RP36-RP37), and mldC (RP38-RP39). Data were

normalized to mRNA levels of the C. difficile housekeeping gene rpoB (primer pair TEQ009-

TEQ010).

Morphology of mldA::erm and mldB::erm mutants

Cultures grown overnight in TY medium were diluted 1:100 into TYN. When the OD600 of

the culture reached ~0.6, 500 μl were transferred to a microfuge tube containing 5 μl of a 500

35

μg/ml stock solution of FM4-64FX (Life Technologies) in DMSO. (This derivative of FM4-64 is

fixable, but we found that fixation caused staining to appear very splotchy, so we omitted the

fixation step.) Samples were then removed from the anaerobic chamber. Cells were collected

by centrifugation, 450 μl of supernatant were removed, and the pellet was suspended in the

residual volume of 50 μl. For microscopy a 2 μl sample was transferred to a thin agarose pad

(Tarry et al., 2009) and imaged using a filter for Texas Red (Chroma # U-N41004).

Complementation experiments were performed similarly, except the overnight cultures

contained 10 μg/ml of thiamphenicol (to select for the plasmid) and the day-cultures contained

both thiamphenicol and tetracycline (to induce plasmid-borne genes). The best

complementation was achieved using tetracycline at 200 ng/ml.

Transmission Electron Microscopy

Wild-type and mldA::erm strains grown overnight in TY medium were diluted 1:100 into

10 mls of TYN. When cultures reached an OD600 of ∼0.7, cells were fixed in the media using

2.5% glutaraldehyde for 30 minutes. Cells were centrifuged at a 5,000 rpm, washed with PBS,

and resuspended in 50 µl of PBS. Warm 2% agarose was added 1:1. Once removed from the

anaerobic chamber, this mixture was allowed to solidify at room temperature and subsequently

at 4˚C for 30 minutes each. Samples were prepared as described previously (Hsiao et al., 2011).

Samples were examined by a JEM-1230 transmission electron microscope (Japan Electron Optics

Laboratory Co., Peabody, MA) at the University of Iowa Central Microscopy Research Facility.

Localization of CFP-Mld fusion proteins in C. difficile

C. difficile strains containing CFP fusion plasmids were grown overnight in BHI containing

thiamphenicol at 10 μg/ml. The next morning, cultures were diluted 1:100 in the same medium

36

supplemented with tetracycline at 500 ng/ml. When cultures reached an OD600 of ∼0.6, a 500 μl

sample was transferred to a microfuge tube containing a mixture of phosphate-buffered saline

(PBS), paraformaldehyde and glutaraldehyde as described (Pogliano et al., 1995). Samples were

then removed from the anaerobic chamber and held at room temperature for 15 minutes,

followed by 30 minutes on ice. Fixed cells were washed 3x with PBS, resuspended in 100 μl of

PBS and left in the dark at room temperature >14 hours to allow chromophore formation. The

cells were immobilized on thin agarose pads for microscopy (Tarry et al., 2009). Our

microscope, camera, and software have been described previously (Mercer and Weiss, 2002).

The filter set for CFPopt was ECFP/DEAC (Chroma # 31044v2).

Localization of FtsZ-GFP in E. coli

Strain EC448 was grown overnight in TY in a Coy anaerobic chamber. The next morning

the cells were diluted 1:500 into TY containing 2.5 μM IPTG (to induce the chromosomal ftsZ-gfp

fusion) and grown to OD600 ~0.6. Cells were then fixed as described for localization of CFP-Mld

fusions in C. difficile. Samples were photographed using a GFP filter set (Chroma no. 41017) at

0, 5, 15, and 20 hours after removal from the anaerobic chamber. To verify that GFP

fluorescence resulted from maturation of protein produced anaerobically, an aliquot of the fixed

cells was spread onto an LB plate and incubated overnight at 37°C. Only a few colonies

appeared, corresponding to 1 CFU per 107 cells in the fixed sample.

Immunoblot Analysis

For immunoblot analysis of CFP fusions, C. difficile strains carrying various CFPopt

plasmids were grown to an OD600 of 0.6-0.8 in 5 mls of BHI supplemented with thiamphenicol

and tetracycline. Cultures were then removed from the anaerobic chamber for further

37

processing. Cells were collected by centrifugation and the resulting pellet was resuspended in

~0.5 mls of Laemmli sample buffer to achieve a sample concentration of 6.0 OD600 units per ml.

Cells were lysed using sets of 10 brief pulses (~0.5 sec each) of sonication (Branson Sonifier 150,

power output 8.5). Lysates were loaded onto an Any kD TM polyacrylamide gel (Bio-Rad), and

proteins were separated by SDS-PAGE. Proteins were then transferred to nitrocellulose. Blots

were blocked with 5% nonfat dry milk in phosphate-buffered saline containing 0.1% Tween

(PBS-T). CFPopt was detected with polyclonal anti-GFP antibody diluted 1:10,000 in blocking

buffer, followed by goat anti-rabbit IgG (H+L) IRDye 800CW obtained from Li-Cor. Finally, blots

were imaged on an Odyssey CLx Infrared Imaging System (Li-Cor).

For immunoblot analysis of toxin production, strains were grown for 5 days in 10 mls of

TY or TY containing 1% glucose. Glucose was included to repress toxin production and served as

a specificity control since we did not have an isogenic tcdA null mutant (Dupuy and Sonenshein,

1998). After removal of cultures from the anaerobic chamber, cells were pelleted by

centrifugation and the resulting supernatant was concentrated using Centricon 100 columns

(Microcon, Millipore, Bedford, MA) to a final volume of 50-100 µl. Concentrated supernatant

was mixed with 6x Laemmli sample buffer proteins and proteins were separated by SDS-PAGE

(Mini-PROTEAN TGX Precast gels from Bio-Rad, 7.5% polyacrylamide). Subsequent steps were

as described above. The primary antibody was Anti-Clostridium difficile toxin A antibody [PCG4]

(ab19953; Abcam, Cambridge, MA) diluted 1:1,000 in PBS and incubated with the blot overnight.

The secondary antibody goat anti-mouse IRDye 800CW (Li-Cor) diluted 1:10,000 in PBS and

incubated for 1 hour.

38

Spore preparation

C. difficile spores for virulence assays were prepared as previously described (Sambol et

al., 2001). Briefly cells were grown on TY plates for 5 days. Plates were then removed from the

anaerobic chamber and allowed to sit in air for at least 2 hours. Cells and spores were

suspended in PBS, washed, and any remaining viable vegetative cells were killed by heat

treatment at 65oC aerobically for 15 minutes. Spore preparations were stored at 4oC until use.

The concentration of viable spores in the preparations was quantified by germination on TY with

0.1% taurocholate. The yield was typically 10 to 25 million spores per plate.

To compare the overall efficiency of sporulation and germination of the wild-type and

mldA::erm mutant strains, spores were prepared as described above using either TY or TYN

plates. Dilutions of these spore preparations were then plated on either TY or TYN, each

containing 0.1% taurocholate, and colony formation was assessed after incubation of the plates

anaerobically for 24 hours at 37°C.

Syrian Hamster Model of C. difficile Infection

For pathogenesis assays, C. difficile spores were prepared as previously described

(Sambol et al., 2001). The number of viable spores was determined by plating on TY containing

0.1% taurocholate and 1% cysteine (Ho and Ellermeier, 2011). Syrian gold hamsters (80-125g,

Harlan Sprague-Dawley, Inc., Indianapolis, IN) were treated with a single oral dose of

clindamycin (30 mg/kg) five days prior to infection with an oral dose of 10,000 spores of either

wild type, mldA::erm mutant or mldB::erm mutant of C. difficile (Lyras et al., 2009; Sambol et al.,

2001). Hamsters were monitored twice daily for signs of morbidity: wet tail, diarrhea, ruffled

fur, and lethargy. Moribund hamsters were euthanized. The data were graphed as Kaplan-

Meier survival analyses (Kaplan and Meier, 1958) and compared for statistical significance using

39

the log-rank test and the software GraphPad Prism 6 (GraphPad Software, San Diego, CA). To

ascertain whether hamsters were colonized with C. difficile, fecal samples were obtained from

each hamster beginning one day prior to clindamycin treatment until day 4 post inoculation.

Fecal samples were homogenized in sterile PBS and serial dilutions plated on CCFA containing

0.1% taurocholate. The animal experiments performed in this study were approved by the

University of Iowa Institutional Animal Care and Use Committee.

Results

Bioinformatic identification of a cell division gene cluster

In many bacteria, proteins containing a SPOR domain are prominent components of the

septal ring that mediates cell division (Arends et al., 2010; Dai et al., 1993; Gerding et al., 2009;

Möll and Thanbichler, 2009). SPOR domains (Pfam 05036) are about 75 a.a. long and bind to the

peptidoglycan sacculus (Brown et al., 2013; Punta et al., 2012; Ursinus et al., 2004). The

sacculus, also called the cell wall, surrounds bacterial cells, giving them their characteristic shape

and protecting them from lysis due to turgor pressure. The genome of C. difficile strain 630

(ribotype 12) encodes a single putative SPOR domain protein, which is designated CD2717 and

has no assigned function (Monot et al., 2011; Punta et al., 2012; Sebaihia et al., 2006). cd2717

sits immediately upstream of two additional genes of unknown function, annotated cd2716 and

cd2715a. As we demonstrate below, all three genes encode midcell localizing division proteins,

so we will henceforth refer to them as MldA, MldB, and MldC (for CD2717, CD2716, and

CD2715a, respectively).

MldA is predicted to be an 847 a.a. protein with a single transmembrane helix, three

coiled-coil domains that might mediate protein-protein interactions, and a SPOR domain near

the C-terminus (Figure 2.1C). The putative SPOR domain in MldA is supported by an E-value of

40

1.009e-8 and a manual alignment of the domain to the Pfam HMM SPOR domain logo [(Punta et

al., 2012); Figure 2.2A]. MldA also has a strikingly high content of charged amino acids—of the

847 residues in MldA, 341 are either D, E, K, or R, corresponding to 40% of the sequence (Figure

2.2B). MldB and MldC are predicted to encode cytosolic proteins of 663 and 106 residues,

respectively (Figure 2.1C). MldB has two predicted coiled-coil regions (Lupas, 1996), but

bioinformatic analyses failed to identify any conserved domains or sequence motifs in MldC.

BLAST searches revealed MldA, MldB, and MldC are found in all sequenced clinical

isolates of C. difficile, and the closely related species C. hiranonis, Peptostreptococcus

anaerobius, and P. stomatis (Table 2.4). Moreover, the order and spacing of the respective

genes are conserved across these organisms, suggesting they comprise a single transcription

unit and encode proteins that function together in the same pathway. Outside of the fact that

SPOR domains are widely conserved, homologs of the Mld proteins could not be identified in

any other sequenced bacteria, even pathogenic clostridia such as C. perfringens, C. botulinum,

and C. tetani. The restricted phylogenetic distribution of the Mld proteins is interesting in light

of a recent proposal to move C. difficile into the family Peptostreptococcaceae (Yutin and

Galperin, 2013). It also raises the possibility that inhibitors of the Mld proteins might target C.

difficile without disrupting the intestinal microbiome.

The mldABC genes are co-transcribed

To determine whether the mld genes are co-transcribed in C. difficile, RNA was

extracted from cells grown to mid-logarithmic phase and RT-PCR was performed using primer

pairs spanning the intergenic regions (Figure 2.1A). Amplification products were obtained

between mldA and mldB, and between mldB and mldC, but not upstream of mldA or

downstream of mldC (Figure 2.1B). These products were not the result of DNA contamination

41

since they were not obtained in the absence of reverse transcriptase (-RT) (Figure 2.1B). As a

positive control for each primer pair, we demonstrated that PCR products could be obtained

with genomic DNA as a template (gDNA) (Figure 2.1B). We conclude that mldA, mldB, and mldC

probably constitute a transcription unit. Moreover, the absence of an RT-PCR amplification

product using the upstream-most primer pair, designated primer pair 1 in Figure 2.1A, implies

the promoter is within about 300 bp of the mldA start codon. This inference follows from the

location of the 5’ primer, which spanned from -328 to -308 with respect to that ATG of mldA and

was too far upstream to capture the 5’ end of the transcript.

mld mutations impair cell division, cell elongation, and cell shape

To determine whether mldA is important for cell division, we used TargeTron

mutagenesis (Heap et al., 2010) to insert a 1.7 kb group II intron (ltrB) carrying an erythromycin

cassette (erm) after nucleotide 248 of the mldA gene (Figure 2.1A). To determine whether mldA

is important for cell division, we used TargeTron mutagenesis (Heap et al., 2010) to insert a 1.7-

kb group II intron (ltrB) carrying an erythromycin cassette (erm) after nucleotide 248 of the mldA

gene (Figure 2.1A). Because the phenotypic changes in morphology resulting from gene mutants

of E. coli and B. subtilis often depend on the concentration and nature of salts in the growth

media (Formstone and Errington, 2005; Murray et al., 1998; Reddy, 2007; Ricard and Hirota,

1973), we examined our mutant in TY medium and TYN medium (TY medium containing 0.5%

NaCl). In TY medium, the mldA::erm mutant and the wild type grew at similar rates, although

examination of cells under the microscope revealed subtle abnormalities in the mutant (Figure

2.3A and B). But when grown in TYN medium, the mutant exhibited a modest reduction in

growth rate and striking morphological defects (Figure 2.3A and C and Table 2.5). These defects

were accentuated by prolonging growth in TYN medium, so to quantify the phenotypic changes,

42

we standardized growth by diluting overnight cultures grown in TY medium 1:100 into TYN

medium and incubating for about 7 h until the OD600 reached 0.6.

The wild-type strain grew as straight rods with an average length of ~9 μm. Staining

with the membrane dye FM4-64 revealed that over 90% of the cells had either zero or one

division septum, indicating daughter cells separate shortly after cytokinesis. In contrast, almost

all of the cells of the mldA::erm mutant had an irregular or curved contour, and ~3% of the cells

exhibited severe bends (white arrow in Figure 2.3). The mldA::erm mutant also exhibited

inefficient separation of daughter cells (a “chaining” phenotype), so that the mutant often

appeared as a filament of 2-8 cells (Figure 2.3C; Table 2.5). On average the cells of the

mldA::erm mutant were 17% shorter than wild type. Prolonged growth in TYN exacerbated

these defects and resulted in some lysis. Transmission electron microscopy suggested lysis is

associated with separation of the peptidoglycan layer from the cell near division septa (Figure

2.3D). Despite the modest lysis seen in TYN broth, the plating efficiency on TYN was the same as

wild-type, although the colonies were smaller. Collectively, these phenotypes indicate MldA is

required for proper elongation and efficient separation of daughter cells at the end of

cytokinesis.

We inactivated mldB by inserting a group II intron immediately after nucleotide 153 of

the gene (Figure 2.1A). The mldB::erm mutant exhibited similar morphological defects to the

mldA::erm mutant, although these were less severe (Figure 2.3; Table 2.5). We were unable to

construct an mldC mutant despite four attempts that targeted different insertion sites. We

doubt that mldC is essential because (polar) insertion mutants in mldA and mldB are viable.

More likely, the potential TargeTron insertion sites in this short (~300 bp) gene are not very

efficient.

43

During the course of these studies, we noticed that C. difficile does not regulate division

site placement very carefully. This can be seen in Figure 2.3B, where one of the wild-type cells

has an eccentric division septum, and in Figure 2.3C, where the distribution of septa in chains of

the mld mutants is sometimes uneven. Division site placement has been studied most

intensively in the rod-shaped bacteria E. coli and B. subtilis, which, in contrast to C. difficile,

divide with high fidelity at the midpoint of the cell (Migocki et al., 2002; Yu and Margolin, 1999).

We do not know the genetic basis or physiological significance of relaxed division site selection

in C. difficile, but it is not an artifact of the JIR8094 strain background because we also observed

irregular constriction site placement in strain R20291.

Complementation of insertions in mldA or mldB requires the entire mldABC operon

Quantitative RT-PCR revealed the erm insertion in mldA is polar on mldB and mldC, but

not the next downstream gene, gtaB (Figure 2.1D). This supports the operon model proposed

above on the basis of intergenic RT-PCR (Figure 2.1B) but also raised the possibility that the

morphology defects seen in the mldA::erm mutant may result from polarity on mldB and/or

mldC. To address this, we first constructed a set of plasmids that contained various

combinations of mldA, mldB, and mldC together with 295 bp of DNA from upstream of mldA,

which we presumed would include the promoter. However, none of these plasmids

complemented the mldA::erm mutant. Specifically, when the entire operon was present on the

plasmid, the cells became filamentous, suggesting overexpression of mldABC was interfering

with cell division. Subsets of genes did not cause filamentation, but they did not rescue any

defects either.

To get better control over gene expression, we cloned the various combinations of

mldA, mldB, and mldC into a plasmid with a tetracycline-inducible promoter (Fagan and

44

Fairweather, 2011). The shape and chaining defects of either the mldA::erm and mldB::erm

mutant backgrounds were largely corrected when all three mld genes were expressed (Figure

2.4 and Table 2.6). Weak complementation was observed with the mldAB construct, but

plasmids expressing mldA, B, or C alone did not affect the phenotypes. Notably,

complementation was very sensitive to the amount of tetracycline used as inducer. We only

observed complementation with 200 ng/ml tetracycline; 100 ng/ml did not correct the

morphological defects, while 400 ng/ml caused cells to become filamentous in some cases (see

below).

We were surprised that expressing mldB and mldC together failed to complement the

mldB::erm mutant. To explore this further, we performed qRT-PCR on the mldB::erm mutant,

which revealed mldA mRNA is 3-fold lower compared to wild type (Figure 2.5). Decreased

expression of mldA in the mldB::erm mutant may be due to altered transcript stability. In sum,

proper expression of mldA, mldB, and mldC was required for complementation, suggesting all

three genes contribute to cell shape and division.

Overproduction of MldAB impairs cell division

During the course of the complementation experiments, we observed a striking

filamentation phenotype when plasmids carrying either mldABC or mldAB were induced with

high levels of tetracycline (Figure 2.4D). For example, when an overnight culture grown in TY

was diluted 1:100 into TYN with 500 ng/ml tetracycline, some filaments were over 100 μm long

when the culture reached an OD600 = 0.5, and with prolonged growth could reach lengths of 1

mm. Staining with FM4-64 revealed very few septa in these filaments, indicating division was

impaired prior to constriction. This defect fundamentally different from the late stage (chaining)

defect observed in mld mutants (Figure 2.3). We do not know why overproduction of MldABC

45

impairs division, but potential mechanisms include sequestration of an essential division protein

or impaired access to the peptidoglycan. A similar filamentation phenotype has been reported

in E. coli when a SPOR domain protein named DamX is overproduced (Arends et al., 2010;

Lyngstadaas et al., 1995). Induction of individual mld genes with high levels of tetracycline did

not result in elongation.

The Mld proteins localize to the midcell

We next wanted to ascertain whether the Mld proteins are components of the septal

ring structure that mediates cytokinesis in bacteria. In most bacteria this question would be

addressed by constructing fusions to GFP, but C. difficile is a strict anaerobe and GFP requires O2

for chromophore maturation. A potential solution to this problem was foreshadowed in an

early paper on GFP by Tsien and colleagues, who demonstrated that GFP produced from a

plasmid in E. coli growing anaerobically became fluorescent over a period of several hours once

air was admitted to the system (Heim et al., 1994). However, we considered it unlikely that

simply expressing a GFP fusion in C. difficile and then exposing the bacteria to air would reveal

septal localization because the bacterial division apparatus is highly dynamic. For example, the

FtsZ ring turns over with a half-life measured in seconds (Anderson et al., 2004), suggesting

septal ring proteins would delocalize if a strict anaerobe were exposed to air for several hours to

allow GFP to mature. These considerations prompted us to ask whether GFP could still acquire

fluorescence if cells grown anaerobically were fixed with crosslinking agents. Indeed, a pilot

experiment using E. coli engineered to produce FtsZ-GFP from a single-copy chromosomal ftsZ-

gfp allele revealed that cells grown and fixed anaerobically were nonfluorescent at first, but the

FtsZ ring became visible after exposure to air overnight (Figure 2.6A).

46

To extend this approach to C. difficile, whose genome has a G+C content of only 29%,

we obtained a derivative of Cyan Fluorescent Protein called “CFPopt” that was codon optimized

for expression in bacteria with AT-rich genomes (Sastalla et al., 2009; Sebaihia et al., 2006). We

chose CFP because the intrinsic fluorescence of C. difficile interferes with visualization of yellow

and green fluorescent proteins. A plasmid that produces CFP-MldA under control of a

tetracycline-inducible promoter was introduced into wild-type C. difficile by conjugation. Cells

were grown to OD600 of ~0.5, fixed with paraformaldehyde/glutaraldehyde, removed from the

anaerobic chamber, transferred to phosphate-buffered saline (PBS), and left at room

temperature in the dark overnight to allow for chromophore maturation. When examined the

next morning, ~34% of the cells in the population contained a single fluorescent band, indicating

CFP-MldA localized to the septal ring (Figure 2.6B; Table 2.7). No other prominent localization

patterns were apparent. We observed similar frequencies of midcell localization for both CFP-

MldB and CFP-MldC (Figure 2.6B; Table 2.7). If CFPopt was not fused to anything, fluorescence

was distributed uniformly throughout the cytoplasm (Figure 2.6B). If the fixation step was

omitted, cells lysed during overnight incubation in aerobic PBS.

To determine whether MldA, MldB, and MldC exhibited any interdependencies for

septal localization, we introduced the CFP-Mld fusion constructs into the mldA::erm and

mldB::erm mutants. Western blotting verified that each fusion was produced at similar levels in

wild-type and mutant backgrounds (Figure 2.7). CFP-MldA localized to the midcell slightly better

in the mutants than in wild type (Table 2.7), indicating MldA does not require either MldB or

MldC for septal localization. In contrast, CFP-MldB failed to localize to the midcell in the

mldA::erm mutant and localized only infrequently (~4%) in the mldB::erm mutant (Table 2.7),

which, as noted above, probably has reduced expression of mldA. Similarly, CFP-MldC failed to

localize in either mutant (Table 2.7). Thus, localization of MldB and MldC to the midcell clearly

47

requires the presence of MldA, and may require the presence of all three Mld proteins. It is not

obvious why CFP-MldA localizes in a higher fraction of mld mutant cells than wild type, but if the

division septum matures slowly in the mutants, the fraction of cells that are late in the division

cycle might increase.

mld mutants are attenuated for pathogenesis in a hamster model of C. difficile infection

We used the Syrian hamster model of C. difficile disease to assess the effect of mld

mutations on pathogenesis (Sambol et al., 2001). Hamsters were given a single oral dose of

clindamycin to disrupt the intestinal microbiome. Five days later, each hamster received an oral

dose of 10,000 C. difficile spores (as assessed by colony-formation), and the animals were

monitored for 30 days for signs of illness. Fecal pellets were collected to determine the

presence of C. difficile spores, an indicator of colonization. The results are summarized in Figure

2.8. Of the 18 hamsters inoculated with wild-type spores, all were colonized. Seventeen

hamsters developed diarrhea and 15 hamsters ultimately required euthanasia. Of the 20

hamsters that received mldA::erm spores, 19 were colonized, 4 developed diarrhea, and 3 were

euthanized. Ten hamsters received mldB::erm spores. Of these, 9 were colonized, 3 developed

diarrhea, and 1 was euthanized.

We did not attempt to complement the virulence defect because of issues related to

plasmid instability and the requirement for a precise concentration of tetracycline to induce

gene expression. However, the fact that we obtained similar results with two completely

independent mutants (mldA::erm and mldB::erm) argues strongly that attenuation is due to loss

of mld function rather than an unknown secondary mutation elsewhere in the genome.

To begin exploring potential explanations for the pathogenesis defect, we assayed

sporulation, germination, and production of toxin A. No defects were found (Figure 2.9 and

48

2.10). It is important to note, however, that these results come with an important caveat—the

phenotypic changes in mld mutants are sensitive to the growth conditions, and it is likely that

the media we use in the laboratory (TY, TYN) do not faithfully reflect conditions in the intestinal

tract of a hamster.

Discussion

Multiple observations argue that the mldABC gene cluster contributes to proper cell

division and elongation in C. difficile. First, MldA is predicted to contain a small, peptidoglycan-

binding domain known as a SPOR domain. Proteins that contain a SPOR domain have been

implicated in cell division in a variety of bacteria (Arends et al., 2010; Dai et al., 1993; Gerding et

al., 2009; Möll and Thanbichler, 2009). The only exceptions of which we are aware concern

proteins involved in other aspects of morphogenesis, namely, swarmer cell differentiation, or

sporulation (Gode-Potratz et al., 2011; Mishima et al., 2005). Second, mldA::erm and mldB::erm

null mutants have a late-stage division defect known as a chaining phenotype. Third,

overexpression of mldABC inhibits an early stage of cell division. Fourth, the Mld proteins

localize to the division site, implying they are part of the apparatus that mediates cell division.

Fifth, cells in chains of the mldA::erm and mldB::erm mutants are short, indicating they do not

elongate very efficiently. Finally, these cells usually have a curved or irregular contour,

indicating they have lost control of lateral wall growth.

A chaining phenotype can result from either slow constriction or delayed separation of

daughter cells. Because the Mld proteins contribute to both division and elongation, we cannot

be certain of where they act in the division process. Nevertheless, our findings point to a

separation defect because if chaining were a consequence of slow inward growth of the septum,

the cells that make up the chains would probably be longer than normal; instead they are

49

shorter. Moreover, the Mld proteins localized in ~35% of the cells, whereas staining with FM4-

64 revealed septa in ~65% of the cells, suggesting the Mld proteins are needed for a late stage

of cell division (like daughter cell separation) rather than an early stage (like initiation of

constriction).

An important question for the future will be to determine the mechanism by which

MldABC facilitates cell division and proper extension of the lateral wall. In principle, defects in

almost any process associated with biogenesis of the cell envelope could impair growth and

division. Perturbations in teichoic acid metabolism are known to cause shape and division

defects in B. subtilis, Listeria monocytogenes, and S. aureus (reviewed in (Brown et al., 2013)).

Alternatively, C. difficile has a proteinaceous S-layer that is anchored to the peptidoglycan wall

and could plausibly affect elongation and division (Calabi et al., 2001; Fagan and Fairweather,

2014). But several observations suggest the most likely role for MldABC is related to biogenesis

of the peptidoglycan cell wall. MldA is predicted to have a peptidoglycan-binding SPOR domain

and transmission electron microscopy revealed disintegration of the cell wall at sites of

septation in an mldA::erm mutant. Moreover, the shape and chaining defects are reminiscent of

previously reported mutants with defects in peptidoglycan hydrolases (or proteins that govern

peptidoglycan hydrolases) in other bacterial species (Cloud and Dillard, 2004; Dominguez-

Cuevas et al., 2013; Heidrich et al., 2001; Meisner et al., 2013; Sham et al., 2011; Uehara et al.,

2010; Yang et al., 2011). Whatever the biochemical activity of the Mld proteins turns out to be,

we suspect all three proteins function as a complex, as reflected in the localization

dependencies, the requirement for all three genes to complement the mldA::erm and mldB::erm

mutants, and the presence of predicted coiled-coil regions in both MldA and MldB.

Our data indicate that mldA::erm and mldB::erm mutants are severely attenuated in the

Syrian hamster model of C. difficile infection. We can envision several ways in which the Mld

50

proteins of C. difficile might contribute to virulence. Because the Mld proteins affect cell

division and rod morphology, they might make a general contribution to growth and fitness.

Nevertheless, the mldA::erm mutant and, even more so, the mldB::erm mutant colonized

hamsters about as well as the parental wild type C. difficile strain (Figure 2.8A). It is possible

that cell wall abnormalities alter the inflammatory response, attachment, toxin secretion,

sporulation, or germination. One challenge in addressing these ideas is that the phenotypic

changes caused by mld mutations are sensitive to growth conditions (e.g., TY vs. TYN), making it

difficult to extrapolate to the hamster model of infection, where the extent of the

morphological defects is not known. Thus, although the mldA::erm mutant appeared normal in

assays of sporulation, germination, and production of toxin A (Figure 2.9 and 2.10), whether

these processes are impaired during growth in a hamster remains to be determined.

Early studies of GFP chromophore formation demonstrated the protein is not

fluorescent when expressed anaerobically in E. coli, but acquires fluorescence after O2 is

admitted to the system (Heim et al., 1994). This was recently confirmed for an O2-dependent

red fluorescent protein (tdTOMATO) produced in C. difficile (Barra-Carrasco et al., 2013). We

have extended this finding by showing GFP and CFP acquire fluorescence even after fixation

with a mixture of paraformaldehyde and glutaraldehyde. These fixatives work by crosslinking

primary amines, especially lysine residues, which are absent from the chromophore of GFP-

family proteins (Tsien, 1998). The advantage of fixation is that it preserves the proper

localization of proteins after the cells are exposed to air and die. This opens the door for using

GFP and related proteins to study protein localization at relatively high resolution in a variety of

strict anaerobes. Although in the present study we only applied fluorescent proteins to the

problem of determining the subcellular localization of proteins, our approach can probably be

51

adapted to a wide array of studies where anaerobiosis is pertinent, such as exploring the

architecture of biofilms or as a reporter in high-throughput screens for new antibiotics.

52

Table 2.1. Strains used in chapter two.

Strain Genotype and description Reference or Source

E. coli EC448 MC4100 Δ(λattL-lom)::bla lacIq P204::ftsZ-gfp (Weiss et al., 1999)

Omnimax – 2 T1R

F´ [proAB+ lacIq lacZΔM15 Tn10(TetR) Δ(ccdAB)] mcrA Δ(mrr-hsdRMS-mcrBC) Φ80lacZΔM15 Δ(lacZYA-argF) U169 endA1 recA1 supE44 thi-1 gyrA96 relA1 tonA panD.

Invitrogen

HB101 / pRK24

F− mcrB mrr hsdS20(rB

− mB

−) recA13 leuB6 ara-14 proA2 lacY1

galK2 xyl-5 mtl-1 rpsL20.

(Trieu-Cuot et al., 1987)

RAN334 Omnimax / pRAN334 (Ptet::cfpopt cat) This study

RAN357 Omnimax / pRAN357 (Ptet:: cfpopt- MCS cat) This study

RAN410 Omnimax / pRAN410 (Ptet:: cfpopt-mldA cat) This study

RAN460 Omnimax / pRAN460 (Ptet:: cfpopt-mldB cat) This study

RAN461 Omnimax / pRAN461 (Ptet:: cfpopt-mldC cat) This study

C. difficile JIR8094 Spontaneous erythromycin-sensitive derivative of C. difficile strain

630. (O'Connor et al.,

2006) RAN125 JIR8094 mldA248::ltrB::ermB This study

RAN258 JIR8094 mldB153::ltrB::ermB This study

RAN427 JIR8094 / pRAN414 (Ptet::mldABC cat) This study

RAN428 JIR8094 / pRAN415 (Ptet::mldAB cat) This study

RAN429 JIR8094 / pRAN416 (Ptet::mldA cat) This study

RAN430 JIR8094 / pRAN417 (Ptet::mldBC cat) This study

RAN431 JIR8094 / pRAN418 (Ptet::mldB cat) This study

RAN432 JIR8094 / pRAN419 (Ptet::mldC cat) This study

RAN433 JIR8094 / pRPF185 (Ptet::gusA cat) This study

RAN446 RAN125 (mldA::ltrB::ermB) / pRAN414 (Ptet::mldABC cat) This study

RAN447 RAN125 (mldA::ltrB::ermB) / pRAN415 (Ptet::mldAB cat) This study

RAN448 RAN125 (mldA::ltrB::ermB) / pRAN416 (Ptet::mldA cat) This study

RAN449 RAN125 (mldA::ltrB::ermB) / pRAN417 (Ptet::mldBC cat) This study

RAN450 RAN125 (mldA::ltrB::ermB) / pRAN418 (Ptet::mldB cat) This study

RAN451 RAN125 (mldA::ltrB::ermB) / pRAN419 (Ptet::mldC cat) This study

RAN452 RAN125 (mldA::ltrB::ermB) / pRPF185 (Ptet::gusA cat) This study

RAN453 RAN258 (mldB::ltrB::ermB) / pRAN414 (Ptet::mldABC cat) This study

RAN454 RAN258 (mldB::ltrB::ermB) / pRAN415 (Ptet::mldAB cat) This study

RAN455 RAN258 (mldB::ltrB::ermB) / pRAN416 (Ptet::mldA cat) This study

RAN456 RAN258 (mldB::ltrB::ermB) / pRAN417 (Ptet::mldBC cat) This study

RAN457 RAN258 (mldB::ltrB::ermB) / pRAN418 (Ptet::mldB cat) This study

53

RAN458 RAN258 (mldB: ltrB::ermB) / pRAN419 (Ptet::mldC cat) This study

RAN459 RAN258 (mldB: ltrB::ermB) / pRPF185 (Ptet::gusA cat) This study

RAN346 JIR8094 / pRAN334 (Ptet:: cfpopt cat) This study

RAN424 JIR8094 / pRAN410 (Ptet:: cfpopt-mldA cat) This study

RAN462 JIR8094 / pRAN460 (Ptet:: cfpopt-mldB cat) This study

RAN463 JIR8094 / pRAN461 (Ptet:: cfpopt-mldC cat) This study

RAN465 RAN125 (mldA::ltrB::ermB) / pRAN410 (Ptet:: cfpopt-mldA cat) This study

RAN466 RAN125 (mldA::ltrB::ermB) / pRAN460 (Ptet:: cfpopt-mldB cat) This study

RAN467 RAN125 (mldA::ltrB::ermB) / pRAN461 (Ptet:: cfpopt-mldC cat) This study

RAN469 RAN258 (mldB::ltrB::ermB) / pRAN410 (Ptet:: cfpopt-mldA cat) This study

RAN470 RAN258 (mldB::ltrB::ermB) / pRAN460 (Ptet:: cfpopt-mldB cat) This study

RAN471 RAN258 (mldB::ltrB::ermB) / pRAN461 (Ptet:: cfpopt-mldC cat) This study

54

Table 2.2. Oligonucleotide primers used in chapter two.

Oligo Purpose Sequence 5`-3`* CDE914 EBS universal primer

CGAAATTAGAAACTTGCGTTCAGTAAAC

NF1438 Reverse primer for sequencing inserts in pRPF185

GATCCAGCACACTGGCATCTTTTTATTTAGGGATTTCTCAC

P1447 IBS for constructing mldA248::erm

AAAAAAGCTTATAATTATCCTTAGAAAAAGATTTAGTGCGCCCAGATAGGGTG

P1448 EBS1 for constructing mldA248::erm

CAGATTGTACAAATGTGGTGATAACAGATAAGTCGATTTAGATAACTTACCTTTCTTTGT

P1449 EBS2 for constructing mldA248::erm

TGAACGCAAGTTTCTAATTTCGGTTTTTTCTCGATAGAGGAAAGTGTCT

RP1 Verifying insertion in mldA gene GAAGCAAAATATGCAGAGC

RP2 Verifying insertion in mldA gene GCGTCTTCTTCTCTCTGATAGTTCC

RP26 Intergenic PCR between cd2718-mldA TAGGTCGCTTGAGCCTATTGGTGA

RP27 Intergenic PCR between cd2718-mldA CCATTTGCCAATAAGTGGTTCTGT

RP28 Intergenic PCR between mldA-mldB Verifying insertion in mldB gene

GAATCAGCTCCTTCTTTGAGTGC RP29 Intergenic PCR between mldA-mldB

Verifying insertion in mldB gene

AAAGGCAAGTTATTTCCATAGAC RP30 Intergenic PCR between mldB-mldC

GAAAGTGAAAGTGCAAGTCTTGATG

RP31 Intergenic PCR between mldB-mldC ACACTGACATCCTCATCTATATCC

RP56 Intergenic PCR between mldC-gtaB GAGTTCTTAGGATTGGACTCAGGG

RP33 Intergenic PCR between mldC-gtaB GCACAGTAAATTGCATGTCCAAGAC

RP34 qRT-PCR for mldA ACAGAACCACTTATTGGCAAATGG

RP35 qRT-PCR for mldA GAGAGTCTAATCCCTCTTTCATAGCC

RP36 qRT-PCR for mldB CAGATGGGTTTGAGCTTAGTGGCA

RP37 qRT-PCR for mldB

ACACCTGTATCCTCATTAACTTCATCTG

RP200 qRT-PCR for mldC GAGTTCTTAGGATTGGACTCAGG

RP201 qRT-PCR for mldC TCCATCAGAATCTAAACCACTATCT

RP58 qRT-PCR for gtaB CCTGCAATTGAAGAAGCTCCATCTG

RP59 qRT-PCR for gtaB TCTCCACCTTTACCAGGTGTCTGT

TEQ009 qRT-PCR for rpoB AAGAGCTGGATTCGAAGTGCGTGA

TEQ010 qRT-PCR for rpoB ACCGATATTTGGTCCCTCTGGAGT

RP47 Sequencing mldA

GTAGTATTTCAATAGAAGATGATGCTGAAGAAGGGG

RP48 Sequencing mldA

CTTCTTTATTGCTGTCGTACTCTTTATCTGTAG

RP80 Sequencing mldB CGATAGCTTGACTATTCTAAAGTC

RP81 Sequencing mldB CATCTTGCATAAGTTTTTGAACCATAC

RP114 IBS for constructing mldB153::erm

AAAAAAGCTTATAATTATCCTTAAATGACGATACAGTGCGCCCAGATAGGGTG

RP115 EBS1 for constructing mldB153::erm

CAGATTGTACAAATGTGGTGATAACAGATAAGTCGATACAGGTAACTTACCTTTCTTTGT

RP116 EBS2 for constructing mldB153::erm

TGAACGCAAGTTTCTAATTTCGGTTTCATTCCGATAGAGGAAAGTGTCT

RP160 Cloning cfpopt into pRPF185

CCCGGATCCTTACTTATATAATTCATCCATTCC

RP161 Cloning cfpopt into pRPF185

GGGGAGCTCCTGCAGTAAAGGAGAAAATTTTATGGTTTCAAAAGGAGAAGAATTATTTAC

RP164 Cloning mldABC, mldAB and mldA into GGGGAGCTCCTGCAGTAAAGGAGAA

55

pRPF185 AATTTTAAGGTTTTGAGTGAAAATAATATAATAAG

RP165 Cloning mldABC and mldC into pRPF185

CCCGGATCCAATTTCATCTAAGATGCCTTTTATATC

RP166 Forward primer for sequencing inserts in pRPF185

CATTGATAGAGTTATTTGTCAAACTAG RP171 Cloning cfpopt – MCS† into pRPF185

GGCGGATCCGGCGCGCCTCAGCTGTTTAATTAAGTCGACGCATGCGTTCATCTTATATAATTCATCCATTCC

RP178 Cloning cfpopt –mldA fusion AAAGCATGCGTGAAGGTTTTGAGTGAAAATAATATAATAAGAAACACTG

RP179 Cloning cfpopt –mldA fusion TTTGGCGCGCCCTAAGCACCTTCAACTATTTCTTCATAAAGTTTATC

RP184 Cloning mldB into pRPF185

GGGGAGCTCCTGCAGTAAAGGAGAAAATTTTATGTTGAAATTAGGAGAAAAAATCATATA

RP185 Cloning mldC into pRPF185

GGGGAGCTCCTGCAGTAAAGGAGAAAATTTTATGTCAAGTAAATACAATGTTTGGACTTT

RP186 Cloning mldA into pRPF185

CCCGGATCCCTAAGCACCTTCAACTATTTCTTCATAAAG

RP187 Cloning mldB and mldAB into pRPF185

CCCGGATCCTTACTTGACATACTTTATTCCTCCTGTG

RP196 Cloning cfpopt –mldB fusion

AAAGCATGCATGTTGAAATTAGGAGAAAAAATCATATATGAGC

RP197 Cloning cfpopt –mldB fusion

TTTGGCGCGCCTTACTTGACATACTTTATTCCTCCTGTG

RP198 Cloning cfpopt –mldC fusion

AAAGCATGCATGTCAAGTAAATACAATGTTTGGACTTT

RP199 Cloning cfpopt –mldC fusion

TTTGGCGCGCCCTAAATTTCATCTAAGATGCCTTTTATATC

* Restriction sites underlined.

† MCS, multicloning site.

56

Table 2.3. Plasmids used in chapter two.

Plasmid Relevant Features Reference

pRPF185 E. coli-C. difficile shuttle vector with tetracycline-inducible promoter. Ptet::gusA cat CD6ori RP4oriT-traJ pMB1ori

(Fagan and Fairweather,

2011) pCE240 E. coli-C. difficile shuttle vector for creating C. difficile

mutants using TargeTron mutagenesis. ltrB::ermB::RAM ltrA cat pIP404ori pMB1ori RP4oriT

(Ho and Ellermeier, 2011)

pMC123 E. coli-C. difficile shuttle vector. cat bla CD6ori RP4oriT pMB1ori

(McBride and Sonenshein,

2011)

pTHE1037 E. coli-C. difficile shuttle vector. cat CD6ori RP4oriT pMB1ori

This Study

pBL100 E. coli-C. difficile shuttle vector for creating C. difficile mutants using TargeTron mutagenesis. ltrB::ermB::RAM ltrA cat bla CD6ori RP4oriT pMBlori

(Bouillaut et al., 2013)

pSW4-CFPopt Template for PCR of cfpopt (Sastalla et al., 2009)

pDSW1215 pCD240/ mldA248 This study

pRAN101 pTHE1037/ mldA248 ltrB::ermB::RAM ltrA This study

pRAN243 pBL100/ mldB153 This study

pRAN414 pRPF185 Ptet::mldABC This study

pRAN415 pRPF185 Ptet::mldAB This study

pRAN416 pRPF185 Ptet::mldA This study

pRAN417 pRPF185 Ptet::mldBC This study

pRAN418 pRPF185 Ptet::mldB This study

pRAN419 pRPF185 Ptet::mldC This study

pRAN334 pRPF185 Ptet:: cfpopt This study

pRAN357 pRPF185 Ptet:: cfpopt-MCS This study

pRAN410 pRPF185 Ptet:: cfpopt-mldA This study

pRAN460 pRPF185 Ptet:: cfpopt-mldB This study

pRAN461 pRPF185 Ptet:: cfpopt-mldC This study

57

Table 2.4. Sequence identities and E-values of MldABC homologs*

Organism MldA MldB MldC

Clostridium difficile CD196 YP_003215577 (2559)

YP_003215576 (2558) YP_003215575 (2557)

847 a.a. 663 a.a. 106 a.a. E-value 0.0 E-value 0.0 E-value 2e-65

Clostridium difficile Bl1 YP_006199787 (13235)

YP_006199786 (13230) YP_006199785 (13225)

847 a.a. 663 a.a. 106 a.a. E-value 0.0 E-value 0.0 E-value 2e-65

Clostridium difficile R20291

YP_003219085 (2606) YP_003219084 (2605) YP_003219083

(2604) 847 a.a. 663 a.a. 106 a.a. E-value 0.0 E-value 0.0 E-value 2e-65

Clostridium difficile NAP07

EFH14798 (2596)

EFH14797 (2595)

EFH14796 (2594)

844 a.a. 663 a.a. 106 a.a. E-value 0.0 E-value 0.0 E-value 5e-63

Clostridium hiranonis EEA85283 EEA85284 EEA85285 839 a.a. 665 a.a. 104 a.a. E-value 7e-113 E-value 0.0 E-value 1e-6

Peptostreptococcus anaerobius CAG:653-L

EFD05099 EFD05083 HMPREF0631_1876 985 a.a. 660 a.a. 58 a.a. E-value 2e-38 E-value 3e-134 E-value 4e-2

Peptostreptococcus stomatis DSM 17678

WP_007789343 Unannotated ORF Unannotated ORF 947 a.a. 575 a.a. 58 a.a. E-value 7e-38 E-value 2e-108 E-value 2.2e-2

*Determined using BLAST searches (Altschul et al., 2005)

58

Table 2.5. Chaining phenotypes of mld mutants

Strain Total

Cells or filaments*

Overall Length†

(µm)

Cell Length‡

(µm)

Septa per Cell or Filament

0 1 2 3 4-5 ≥6

Wild type 970 8.5 ± 2.8 5.1 31% 61% 6% 1% 0% 0% mldA::erm 908 11.8 ± 4.5 4.3 8% 38% 22% 21% 9% 3% mldB::erm 915 9.6 ± 3.6 4.3 13% 54% 19% 9% 4% 1%

* At least 200 cells (or filaments of cells) of each strain were scored in 4 separate experiments and data were pooled.

† Mean ± standard deviation of the pooled data set. End-to-end length of each cell (or filament of cells) regardless of the number of septa present.

‡ For an isolated cell, this is the distance between the cell poles. For a filament of cells, this the distance between septa. Each cell or filament was measured and divided by the number of septa plus one.

59

Table 2.6. Complementation of mld mutants.

* Mean ± standard deviation, in μm.

Strain Cells Scored

Cell Length*

Septa per Cell host plasmid 0 1 2 3 4-5 ≥6

Wild type vector 451 8.7 ± 2.9 36.8% 59.9% 3.1% 0.2% 0% 0% mldA::erm vector 315 11.7 ± 6.0 13.3% 30.5% 21.9% 14.3% 14.3% 5.7% mldA::erm mldABC 323 10.9 ± 4.0 20.7% 60.7% 14.9% 2.8% 0.9% 0% mldA::erm mldAB 328 13.1 ± 5.6 11.3% 48.8% 20.7% 14.6% 3.7% 0.9% mldA::erm mldA 217 11.6 ± 5.6 11.1% 38.2% 20.7% 15.2% 11.1% 3.7% mldA::erm mldBC 212 11.0 ± 5.6 12.3% 37.3% 17.5% 17.5% 10.4% 5.2% mldA::erm mldB 201 10.8 ± 4.9 12.9% 37.8% 15.4% 13.9% 15.9% 4.0% mldA::erm mldC 219 11.2 ± 5.1 9.1% 39.7% 20.1% 18.3% 9.1% 3.7%

mldB::erm vector 211 13.9 ± 7.0 8.1% 38.4% 14.7% 11.4% 14.7% 12.8% mldB::erm mldABC 207 11.4 ± 5.0 41.1% 53.1% 4.3% 1.4% 0% 0% mldB::erm mldAB 229 14.6 ± 10.4 17.5% 55.0% 14.0% 11.8% 1.7% 0% mldB::erm mldA 109 15.5 ± 9.8 8.3% 31.2% 16.5% 13.8% 20.2% 10.1% mldB::erm mldBC 105 11.3 ± 6.4 12.4% 34.3% 13.3% 21.9% 12.4% 5.7% mldB::erm mldB 107 11.5 ± 8.5 14.0% 27.1% 12.1% 19.6% 15.0% 12.1% mldB::erm mldC 106 11.5 ± 6.6 9.4% 23.6% 16.0% 15.1% 19.8% 16.0%

60

Table 2.7. Dependency of localization on other Mld proteins.

Host CFPopt- MldA

CFPopt-MldB

CFPopt- MldC CFPopt

Wild type 34% 38% 38% 0%* mldA::erm 45% 0%* 0%* n.d.† mldB::erm 64% 4% 0%* n.d.†

*Over 120 cells were scored, so the detection limit is ~1%.

† n.d., not determined.

61

62

Figure 2.1. The mldABC gene cluster. (A) Arrangement of mldABC and neighboring genes. Filled

triangles depict the location of erm insertions used to inactivate mldA and mldB. Arrows above

the triangles show the direction of erm transcription. Numbered lines depict the approximate

locations of products from intergenic RT-PCR. (B) mldABC probably comprise a transcription

unit as determined by intergenic RT-PCR. +RT, reverse transcriptase step included; −RT, reverse

transcriptase step omitted; gDNA, genomic DNA used as the template. (C) Domain organization

of the Mld proteins. For clarity, the predicted domains are not drawn to scale. C, coiled-coil; T,

transmembrane helix; S, SPOR domain; a.a., amino acid. There are no predicted domains in

MldC. (D) Polarity of the mldA::erm mutant as assayed by qRT-PCR.

63

64

Figure 2.2. Sequences of Mld proteins. (A) The putative SPOR domain from MldA (residues 717-

789) was aligned by hand to the Pfam HMM logo for SPOR domains (Schuster-Bockler et al.,

2004). The alignment revealed matches to one of the top three amino acids at 28 out of 76

positions, consistent with previous evidence that SPOR domain sequences are highly degenerate

(Williams et al., 2013). (B) The predicted amino acid sequences of MldA, MldB, and MldC.

Positively charged residues are in blue, negatively charged residues are in red. For MldA, the

predicted coiled-coil regions are underlined in gray, the transmembrane helix is highlighted in

gray, and the SPOR domain is underlined in black. Predicted coiled-coil regions in MldB are

underlined in gray (Lupas, 1996).

65

66

Figure 2.3. Phenotypes of mldA::erm and mldB::erm mutants. (A) Growth curves in TY and TYN.

Each data point represents the mean ± standard deviation (s.d.) of results from triplicate

cultures, but in most cases the error bars are not visible because they are smaller than the

symbols. The data are representative of the results of at least three experiments. (B)

Fluorescence micrographs of TY-grown cells stained with FM4-64. The caret indicates an off-

center division septum in the wild type. (C) Fluorescence micrographs of TYN-grown cells

stained with FM4-64. The white arrow indicates a highly curved segment of an mldA::erm

mutant cell. Similar deformations were observed in 3% of the cells. (D) Transmission electron

micrographs of septal regions from the wild type and an mldA::erm mutant grown in TYN. Black

arrows point to regions where the peptidoglycan appears to be peeling away from the cell in the

mutant. Similar defects were observed in 6 of 22 micrographs of septa from the mldA::erm

mutant but in none of 18 micrographs of septa from the wild type.

67

68

Figure 2.4. Complementation of mldA::erm and mldB::erm. (A) Fluorescence micrographs of

cells stained with FM4-64. Tetracycline was used at 200 ng/ml to induce expression of plasmid-

borne genes. (B) Quantitative summary of the extent of chaining. (C) Overview of

complementation results obtained with different combinations of mld genes. Restoration of

nearly wild-type morphology is indicated by +++, while the absence of complementation is

indicated by −. Quantitative data are presented in Table 2.6. (D) The experiment was

performed as described for panel A, but tetracycline was used at 500 ng/ml.

69

70

Figure 2.5. Expression of mldA is reduced in an mldB::erm mutant. RNA was harvested from

wild-type and mutant C. difficile strains in logarithmic growth, and qRT-PCR was used to assess

mRNA levels for mldA and mldB. The housekeeping gene rpoB was used for normalization.

Oligonucleotide primers are listed in Table 2.2.

71

72

Figure 2.6. Septal localization of GFP and CFPopt fusions to division proteins. (A) Phase and

fluorescence micrographs of E. coli cells producing an FtsZ-GFP fusion. Cells were grown and

fixed anaerobically and then exposed to air for 5 or 20 h, as indicated. (B) Fluorescence

micrographs of wild-type C. difficile cells producing the indicated CFPopt-Mld fusion proteins.

Cells were grown and fixed anaerobically and then exposed to air overnight prior to imaging.

Bright bands near the midcell demonstrate localization of the tagged Mld protein to a division

site. These photographs used an exposure time of 4 s (CFP fusions) or 2 s (CFP alone).

73

74

Figure 2.7. Expression of CFPopt-Mld fusion proteins in wild-type and mld mutant backgrounds.

Whole cell extracts were prepared from the same cultures used for localization (Figure 2.6).

Steady-state levels of CFPopt fusion proteins were determined by Western blotting with

polyclonal anti-GFP antibodies that recognize CFPopt. Note that the same amount of

tetracycline (500 ng/ml) was used for induction in all cases, so the differences in fusion protein

abundance may reflect differential stability. The Western results are consistent with the relative

brightness of the CFP fusions in Figure 2.6. Molecular mass standards are indicated to the left of

the blot. Asterisks denote the various CFPopt fusion proteins, the predicted masses of which

are CFPopt-MldA = 126 kDa, CFPopt-MldB = 106 kDa, CFPopt-MldC = 39.5 kDa, CFPopt = 27.7

kDa.

75

76

Figure 2.8. Mutants of mldA and mldB are attenuated in hamsters. Syrian hamsters inoculated

orally with 10,000 viable spores were monitored for colonization for up to 4 days and morbidity

for up to 30 days. (A) Time from inoculation to detection of C. difficile spores in fecal pellets.

Each point represents one hamster, and the bar represents the mean. Colonization rates were

18/18 for the wild type (WT), 19/20 for the mldA::erm strain, and 9/10 for the mldB::erm strain.

Data from the two hamsters that were not colonized were excluded from calculations of means.

(B) Time from colonization to death. The surviving hamsters were sacrificed on day 30, and their

data were included in calculations of means. (C) Kaplan-Meier survival plot indicating time from

inoculation to death. Numbers in parentheses refer to the number of hamsters in each

treatment group. Data for the wild-type and mldA::erm strains were pooled from two

experiments; the mldB::erm mutant was assayed once. *, P ≤ 0.0003 for comparisons of data

from the respective mutants and the wild type, as determined by logrank test in GraphPad

prism.

77

78

Figure 2.9. Sporulation and germination. Wild type and mutant C. difficile strains were allowed

to sporulate on TY or TYN plates. Spores were harvested using equal volumes of PBS and then

heat-killed at 65°C for 10 min. Serial dilutions were spread onto either TY or TYN containing

0.1% taurocholate to stimulate germination of spores. No colonies were detected on plates that

lacked taurocholate. Samples were normalized to volumes of PBS used to process each plate.

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80

Figure 2.10. Toxin A production. Cell-free spent medium was concentrated approximately 100-

fold and analyzed by Western blotting with a monoclonal antibody against toxin A (predicted

mass 306 kDa). Molecular mass standards are indicated to the left of the blot. Glucose in the

growth medium is known to repress toxin A production and was included in some cultures to

verify antibody specificity (Dupuy and Sonenshein, 1998). Samples were normalized to volume

of spent medium.

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CHAPTER III

Use of mCherry Red Fluorescent Protein for Studies of Protein Localization and Gene Expression in Clostridium difficile

Introduction

Clostridium difficile is a low-GC, spore-forming bacterium that is burdening the health

care systems of developed countries (CDC, 2013; Jones et al., 2013; Rupnik et al., 2009). While

genetic techniques to study C. difficile are becoming increasingly available, the repertoire of

tools remains limited. This is due in part to the strictly anaerobic environment required to grow

C. difficile.

Green fluorescent protein (GFP) can be produced in cells grown anaerobically, but it is

unable to fluoresce because chromophore maturation requires O2 for dehydration reactions

that introduce double bonds into amino acids (Heim et al., 1994). Nevertheless, GFP produced

in an anaerobic environment can mature and fluoresce upon subsequent exposure to air (Heim

et al., 1994; Zhang et al., 2005). We recently took advantage of this observation to show that

GFP can be used to localize cell division proteins in anaerobically grown Escherichia coli (Ransom

et al., 2014). Similarly, we showed that a derivative of cyan fluorescent protein named CFPopt

(because it has been codon optimized for low-GC bacteria) can be used to localize cell division

proteins in anaerobically grown C. difficile (Ransom et al., 2014). In both organisms, it was

necessary to fix cells anaerobically to preserve their architecture and then expose them to air

overnight to allow chromophore maturation, which required many hours. Fixation was

necessary in the case of E. coli to ensure that the localization observed reflected anaerobic

conditions rather than subsequent adaptation to air. In the case of C. difficile, fixation was

necessary because cells lyse soon after transfer to air. Moreover, O2 poisons energy metabolism

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in C. difficile and the divisome disassembles quickly (<2 min) when cells become depleted of

energy (Anderson et al., 2004; Rueda et al., 2003; Strahl and Hamoen, 2010).

One limitation of GFP and CFP for work in C. difficile is that the organism has

considerable intrinsic green and blue autofluorescence. In contrast, there is virtually no red

autofluorescence. We characterize here a codon-optimized allele of the gene encoding red

fluorescent protein mCherry that we call “mCherryOpt.” An alternative red fluorescent protein,

tdTOMATO, has been used with limited success as a reporter in C. difficile; however,

requirements for its use have not been extensively investigated (Barra-Carrasco et al., 2013).

We focused our efforts on mCherry because it is half the size and folds much faster than

tdTOMATO, although it is reported to be less bright (Shaner et al., 2004). We show here that

mCherryOpt produced in C. difficile is fully fluorescent within 2 h of exposure to air and that

interference from intrinsic background fluorescence is negligible. We also describe plasmids

that facilitate using mCherryOpt as a reporter of protein localization and gene expression in C.

difficile. We hope that the availability of these plasmids and mCherryOpt will promote studies

of the basic biology of C. difficile and other low-GC Gram-positive bacteria.

Materials and Methods

Strains, media, and growth conditions

Bacterial strains are listed in Table 3.1. All C. difficile strains are derived from the

erythromycin-sensitive JIR8094 isolate, which is in turn derived from the strain CD630 (O'Connor

et al., 2006; Sebaihia et al., 2006). E. coli OmniMAX 2 T1R and XL1-Blue were used for cloning,

and HB101/pRK24 was used for conjugations. Tryptone yeast extract (TY) medium consisted of

3% tryptone, 2% yeast extract, and 0.1% L-cysteine, plus 2% agar for plates. Luria-Bertani (LB)

medium contained 10% tryptone, 5% yeast extract, and 1% NaCl, plus 1.5% agar for plates. C.

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difficile was grown in TY medium supplemented as needed with thiamphenicol (Thi) at 10 μg/ml,

kanamycin at 50 μg/ml, or cefoxitin at 16 μg/ml. Genes under the control of the Ptet promoter

were induced with anhydrotetracycline hydrochloride (ATc; Sigma, St. Louis, MO). C. difficile

strains were maintained at 37°C in an anaerobic chamber (Coy Laboratory Products, Grass Lake,

MI) in an atmosphere of 10% H2, 5% CO2, and 85% N2. E. coli strains were grown at 37°C in LB

medium supplemented as needed with ampicillin at 200 μg/ml or chloramphenicol at 20 μg/ml.

Plasmid and strain construction

The oligonucleotide primers used in the present study are listed in Table 3.2 and were

synthesized by Integrated DNA Technologies (Coralville, IA). All plasmids are listed in Table 3.3.

Newly constructed plasmids for the present study were derived from pRPF185, which has a

tetracycline-inducible promoter upstream of the gusA gene (Fagan and Fairweather, 2011). To

replace gusA with gfpmut2, primers RP157 and RP158 were used to amplify gfpmut2 from

pGFPmut2 (Cormack et al., 1996) by PCR. The resulting PCR product was digested with SacI and

BamHI and ligated into the same sites of pRPF185 to generate pRAN332. mCherryOpt was

synthesized by GeneArt (Life Technologies, Grand Island, NY) and delivered in a high-copy-

number plasmid named pMA-T-mCherryOpt. This plasmid was digested with SacI and BamHI,

and the 735-bp fragment encoding mCherryOpt was ligated into SacI/BamHI-digested pRPF185

to generate pDSW1728. To create a derivative with an in-frame multiple-cloning site (MCS)

suitable for making gene fusions, mCherryOpt was amplified by PCR with pDSW1728 as the

template and primers RP204 and RP203, the latter of which encodes the MCS. The resulting PCR

product was digested with SacI and BamHI and then ligated into the same sites of pRPF185 to

create pRAN473. PCR was used to amplify a zapA homolog (cd0701) from the JIR8094

chromosome with primers RP176 and RP177. The resulting PCR fragment was digested with

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SphI and AscI and then ligated into pRAN357 (Ransom et al., 2014) to create pRAN409

(Ptet::cfpopt-zapACd). Digestion of pRAN409 with SphI and AscI yielded a zapA restriction

fragment that was ligated into SphI/AscI-digested pRAN473 to create pRAN534

(Ptet::mCherryOpt-zapACd). To create pRAN535 (Ptet::mCherryOpt-mldA), mldA was moved as a

SphI/AscI restriction fragment from pRAN410 (Ptet::cfpopt-mldA) (Ransom et al., 2014) into

pRAN473. To construct pRAN738, the pdaV promoter was amplified from JIR8094 chromosomal

DNA using the primers RP306 and RP307. The PCR product was digested with NheI and SacI and

ligated into pRPF185 digested with the same enzymes. The resulting plasmid carries sequences

extending from positions −320 to −29 with respect to the A of the ATG start codon of pdaV. All

plasmids were introduced into C. difficile strains by conjugation from strain HB101/pRK24 and

selecting for thiamphenicol resistance (Heap et al., 2010; Ho and Ellermeier, 2011).

Bioinformatics

Codon usage was analyzed with the Codon Usage Database at

http://www.kazusa.or.jp/codon/cgi-bin/showcodon.cgi?species=272563 (Nakamura et al.,

2000). ZapA sequences were aligned using Clustal Omega (Sievers et al., 2011).

Comparison of mCherryOpt, CFPopt, and GFPmut2

To compare different fluorescent proteins, C. difficile JIR8094 containing various

plasmids was grown overnight in TY with 10 μg of Thi/ml. The next morning, these cultures

were diluted 1:100 into the same medium containing ATc at 400 ng/ml to induce expression of

plasmid-borne genes. Cultures were grown to an optical density at 600 nm (OD600) of 0.6, and

then cells from 500 μl of culture were fixed as described below, washed, suspended in 30 μl of

phosphate-buffered saline (PBS), and processed for microscopy as described previously

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(Pogliano et al., 1995; Ransom et al., 2014). For convenience, fluorescence was typically allowed

to develop during an overnight (∼16 h) exposure to air, although much shorter incubations are

sufficient for mCherryOpt.

Kinetics of fluorescence aquisition

To determine the rate of mCherryOpt maturation, cultures were grown and processed

as described above except that, as a control, an aliquot of cells was processed without fixation.

Fluorescence was monitored by microscopy or using an Infinite M200 Pro plate reader (Tecan,

Research Triangle Park, NC). To monitor fluorescence in the plate reader, 20 μl of cells in PBS

were added to 180 μl of PBS and transferred to the well of a flat-bottom 96-well microtiter plate

(AS Plate-PS-96-F-C; AG Advangene, IL). Fluorescence was recorded at 15-min intervals

(excitation, 554 nm; emission, 610 nm; gain setting, 100) with a 10-s shake and a 20-s pause

immediately prior to each reading. The cell density (OD600) was recorded at the start and end of

the experiment to correct for cell number but not more often in order to minimize the bleaching

of mCherryOpt.

Protein localization

For studies of protein localization, we recommend inducing at a variety of ATc

concentrations for a variety of times because each fusion protein is unique. We observed

convincing septal localization of mCherryOpt fusions to MldA and ZapA using the simple

induction protocol described above wherein an overnight culture is diluted 1:100 into TY

containing ATc and grown to mid-log phase before fixation. However, the micrographs shown in

the present study were obtained by diluting overnight cultures 1:100 into TY containing a low

level of ATc (20 ng/ml) and grown to an OD600 of 0.3. A higher concentration of ATc (80 to 160

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ng/ml; it varied from day to day) was then added to boost the expression of plasmid-borne gene

fusions. After 1 h, the cells were fixed and processed for microscopy as described below and

elsewhere (Ransom et al., 2014).

Quantifying expression of PpdaV::mCherryOpt

To monitor the induction of PpdaV::mCherryOpt by lysozyme, overnight cultures grown in

TY containing 10 μg of Thi/ml were diluted 1:100 into the same medium. To determine the

effect of a 4-h induction, lysozyme (or water, as a control) was added when the OD600 reached

0.05, and incubation was continued for 4 h until the OD600 reached 0.8. To determine the effect

of a 1-h induction, lysozyme (or water) was added at an OD600 of 0.4, and incubation was

continued for 1 h until the OD600 reached 0.8. An aliquot of each culture was transferred to

ethanol-acetone, and the mRNA levels were determined by quantitative reverse transcription-

PCR (qRT-PCR) as described previously (Ho and Ellermeier, 2011; Ransom et al., 2014) using

Superscript III and Sybr green master mix (Life Technologies, Carlsbad, CA) with the following

gene-specific primer pairs: rpoB, TEQ009 and TEQ010; pdaV, RP376 and RP377; and

mCherryOpt, RP378 and RP379. A second aliquot of each culture was fixed with

paraformaldehyde-glutaraldehyde, and mCherryOpt fluorescence was measured using a Tecan

plate reader as described above.

Fixation protocol

Cells were fixed for microscopy by adapting procedures developed for Bacillus subtilis

(Pogliano et al., 1995). A 16% (wt/vol) paraformaldehyde aqueous solution (methanol-free;

catalog no. AA433689M) was obtained from Alfa Aesar (Ward Hill, MA). A 25% glutaraldehyde

aqueous solution (catalog no. 16220) was obtained from Electron Microscopy Services (Hatfield,

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PA). A 5× fixation cocktail was prepared fresh each day and consisted (per sample) of 20 μl of 1

M NaPO4 buffer (pH 7.4), 100 μl of 16% paraformaldehyde, and 4 μl of 25% glutaraldehyde. The

cocktail was transferred into the anaerobic chamber immediately prior to use, and 120-μl

aliquots were dispensed to 1.5-ml microcentrifuge tubes that had been in the chamber for at

least 24 h. A 500-μl aliquot of cells (OD600 ∼ 0.6) in growth medium was added directly to a

microcentrifuge tube containing 120 μl of the fixation cocktail, mixed by pipetting up and down

three times, and then allowed to sit for 30 min, at which time the sample was removed from the

chamber and incubated on ice for 60 min. Fixed cells were washed three times with PBS,

resuspended in 30 μl of PBS, and left in the dark at room temperature to allow chromophore

maturation. Because the fixation step was typically not completed until late in the afternoon,

fluorescence microscopy was usually not performed until the next morning but, as shown

below, exposure to air for 2 h appears to be sufficient for mCherryOpt to reach maximum

fluorescence. Shorter fixation times (e.g., 15 min at room temperature plus 30 min on ice) and

removal of the samples from the anaerobic chamber immediately after mixing with the fixation

cocktail yielded similar results (not shown).

Microscopy

For imaging, the cells were immobilized on thin agarose pads. Phase-contrast and

fluorescence micrographs were recorded on an Olympus BX60 microscope equipped with a

×100 UPlanApo objective (numerical aperture, 1.35). Images were captured using a black-and-

white Spot 2-cooled charge-coupled device camera (Diagnostic Instruments, Sterling Heights,

MI) with a KAF1400E chip (class 2), a Uniblitz shutter, and a personal computer with Image-Pro

software version 4.1 (Media Cybernetics, Silver Spring, MD). Filter sets for fluorescence imaging

were from Chroma Technology Corp. (Brattleboro, VT). The GFP filter set (catalog no. 41017)

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comprised a 450- to 490-nm excitation filter, a 495-nm dichroic mirror (long pass), and a 500- to

550-nm emission filter. The CFPopt filter (catalog no. 31044v2) comprised a 426- to 446-nm

excitation filter, a 455-nm dichroic mirror (long pass), and a 460- to 500-nm emission filter (band

pass). For mCherryOpt the filter set (catalog no. 41004) comprised a 538- to 582-nm excitation

filter, a 595-nm dichroic mirror (long pass), and a 582- to 682-nm emission filter.

Fluorescence micrographs were captured using 3-s exposure times and identical display

range settings in all cases, so the images can be compared directly. Black level subtraction and

chip defect correction were “on”; all other processing options were “off.” To ensure

comparability of fluorescence images, the display range option was adjusted identically for all

images. Micrographs were cropped, and figures were assembled in Adobe Illustrator (Adobe

Systems, Inc., San Jose, CA).

Plasmid Copy Number

Real-time qRT-PCR experiments were performed as previously described (Ho and

Ellermeier, 2011) using Sybr green master mix (Applied Biosystems) and the following gene-

specific primer pairs: rpoB, TEQ009 and TEQ010; catR, RP314 and RP315; and mldA, RP316 and

RP317. C. difficile chromosomal DNA was harvested from cells grown to mid-log phase. Cells

were lysed as described previously (Ho and Ellermeier, 2011), except that the lysozyme step was

omitted. The DNA was extracted using a phenol-chloroform protocol (Bouillaut et al., 2011).

Plasmid Stability

Plasmid maintenance was determined using a protocol similar to that described by Heap

et al. (Heap et al., 2009). Briefly, JIR8094 containing various plasmids was grown overnight in TY

medium with Thi at 10 μg/ml. Residual Thi was removed by pelleting cells from 1 ml of the

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culture and suspending the pellet in 1 ml of TY medium without antibiotic. Washed cells were

diluted 1:100 into TY medium without antibiotic every 12 h for 3.5 days, for a total of seven

passages, at which point dilutions of the cultures were plated on TY without antibiotic. At least

50 colonies were patched onto TY medium ± 10 μg Thi/ml to determine the fraction of cells that

contained the plasmid. Strain JIR8094 (This) was patched as a negative control. Assuming

cultures return to maximum cell density in 12 h, there are 6.64 doublings per 12-h growth

period (i.e., 26.64 = 100), which corresponds to 46 generations after seven passages.

Segregational stability per generation was calculated using R(1/n), where n is the number of

generations, and R is the fraction of ThiR colonies.

Nucleotide sequence accession number

The DNA sequence of codon-optimized mCherry (mCherryOpt) has been submitted to

GenBank and is available under the accession number KM983420.

Results

mCherryOpt is superior to CFP in C. difficile

Many factors go into determining which fluorescent protein is “best” for any particular

use (Shaner et al., 2005). The monomeric red fluorescent protein mCherry has a generally

favorable combination of brightness, photostability, and rapid maturation, but it has a relatively

high GC content because it has been codon optimized for expression in mammalian cells (Shaner

et al., 2004; Shaner et al., 2005). The GC content of the C. difficile 630 chromosome is only 29%

(13). Analysis of the DNA sequence of mCherry revealed that it is 62% GC and that >90% of the

codons are not commonly used in endogenous C. difficile genes (Table 3.4). Sastalla et al.

showed that production of fluorescent proteins in low-GC Gram-positive bacteria is greatly

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improved by modifying the genes to make them more AT-rich (Sastalla et al., 2009). We

therefore designed a synthetic mCherry gene that was codon optimized for expression in C.

difficile (Figure 3.1). The modified gene, which we will refer to as mCherryOpt, is 30% GC, and

only 1% of the codons are unfavorable for C. difficile (Table 3.4).

We constructed pDSW1728 by cloning mCherryOpt into pRPF185 (Fagan and

Fairweather, 2011), an E. coli-C. difficile shuttle vector with a tetracycline-inducible promoter

(Figure 3.2). Exconjugants of C. difficile were bright red when grown in TY medium containing

anhydrotetracycline (ATc) and visualized by fluorescence microscopy (Figure 3.3). Half-maximal

fluorescence intensity was obtained after ∼1 h of exposure to air and did not require fixation

(Figure 3.4). However, there was extensive lysis in the unfixed samples (data not shown), so

fixation is necessary for studies that require preservation of cellular architecture. Importantly,

control cells carrying the empty vector pRPF185 had essentially no detectable fluorescence in

the red channel. (The bright spots visible in Figure 3.3D come from debris and were useful for

aligning the fluorescence and phase images.) Although C. difficile strains producing CFPopt or

GFPmut2 were fluorescent in the blue and green channels, respectively, the corresponding

pRPF185 controls had considerable background fluorescence, particularly in the green channel.

Thus, at least in C. difficile, mCherryOpt has a significantly better signal-to-noise ratio than the

other two fluorescent proteins.

Application of mCherryOpt to protein localization

We constructed pRAN473 by introducing an in-frame MCS at the 3′ end of mCherryOpt

to facilitate fusing mCherryOpt to the N-terminus of proteins of interest (Figure 3.2). As proof of

principle, we fused mCherryOpt to MldA (locus tag CD2717), a recently described cell division

protein found only in C. difficile and a few of its closest relatives (Ransom et al., 2014). We

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reported previously that a CFPopt-MldA fusion localized to the division site in ∼35% of cells

growing in TY (Ransom et al., 2014). Similar results were obtained with mCherryOpt-MldA, with

septal localization apparent in 41% ± 3% of the population (mean ± the standard deviation, n =

three independent experiments with >100 cells scored per experiment) (Figure 3.5).

Next, we sought to determine whether mCherryOpt could be used to verify the identity

of a predicted C. difficile cell division protein. ZapA is a widely conserved FtsZ-binding protein

that has been studied primarily in E. coli and B. subtilis, where it localizes sharply to the septal

ring (Buss et al., 2013; Galli and Gerdes, 2010; Gueiros-Filho and Losick, 2002; Johnson et al.,

2004; Scheffers, 2008; Small et al., 2007). ZapA proteins have two domains, a globular head that

binds FtsZ and a C-terminal coiled-coil that mediates tetramerization (Low et al., 2004; Roach et

al., 2014). In C. difficile 630, locus CD0701 is annotated as a “putative cell division protein”

based on its similarity to ZapA (Monot et al., 2011; Sebaihia et al., 2006), and we will refer to it

henceforth as ZapACd. However, whether ZapACd is in fact a ZapA ortholog is not completely

clear based on bioinformatic approaches alone; ZapACd is ∼100 amino acids longer than ZapAEc

or ZapABs, and amino acid identity in the region of overlap is modest, e.g., 14% identity to ZapAEc

and 33% identity to ZapABs (Figure 3.6) (Sievers et al., 2011). Nevertheless, we found that an

mCherryOpt-ZapACd fusion protein localized near the midcell in 38% ± 3% of C. difficile cells (n =

three independent experiments with >100 cells scored per experiment) (Figure 3.5). Because

not all mCherryOpt-ZapACd bands were precisely at the midcell, we sought independent

evidence that they correspond to division sites. Measurements of the distance from the cell

poles to the bands revealed most are at or near the midcell, and their spatial distribution mimics

division septa as determined by staining with the membrane dye FM4-64 (Figure 3.7). These

findings are consistent with the prediction that CD0701 is a ZapA ortholog. The anomaly of

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ZapACd being ∼100 amino acids longer than ZapAEc or ZapABs can be attributed to a much longer

predicted coiled-coil domain [Pfam version 27.0 (Finn et al., 2014)].

Application of mCherryOpt as a reporter of gene expression

In C. difficile the pdaV promoter (PpdaV) directs transcription of a seven-gene operon

necessary for resistance to lysozyme (Ho and Ellermeier, 2011; Ho et al., 2014). One of the

genes in the operon is csfV, which codes for an extracytoplasmic function (ECF) sigma factor

required for transcription from PpdaV (Ho et al., 2014). To test the suitability of mCherryOpt as a

gene expression reporter in C. difficile, we replaced the Ptet promoter-regulatory region in

pDSW1728 with a ∼300-bp DNA fragment carrying the promoter-regulatory region for the pdaV

operon. The resulting PpdaV::mCherryOpt reporter plasmid is designated pRAN738.

C. difficile exconjugants harboring pRAN738 were grown to mid-log phase in TY medium,

exposed to lysozyme for 1 h, and then fixed and exposed to air overnight to allow for

chromophore maturation. Fluorescence microscopy revealed the expected lysozyme-

dependent increase in fluorescence intensity, which was completely dependent upon the ECF

sigma CsfV (Figure 3.8A and B). Fluorescence was relatively uniform from cell to cell, indicating

the plasmid is well maintained and that lysozyme reaches all cells in the population. The fact

that the wild-type reporter strain exhibited faint fluorescence even in the absence of lysozyme,

whereas the csfV mutant background did not, confirms basal transcription at PpdaV requires σV

(17, 35). Quantifying fluorescence with a plate reader revealed that there was a 9.4-fold

induction of PpdaV::mCherryOpt (Figure 3.8C; 0 versus 20 μg of lysozyme/ml).

Fluorescent proteins require considerable time to fold and mature, so mCherryOpt

might under-report induction if a significant fraction of the protein fails to become fluorescent.

We explored this possibility by using both qRT-PCR and fluorescence to measure gene

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expression after cells had been exposed to lysozyme for 1 or 4 h. At the 1-h time point,

PpdaV::mCherryOpt was induced 5-fold as measured by fluorescence but 28-fold as measured by

qRT-PCR (Figure 3.9). This is a ∼6-fold difference. At the 4-h time point, the two methods gave

more similar results: 45-fold induction by qRT-PCR and 30-fold induction by fluorescence (Figure

3.9). Collectively, these data indicate mCherryOpt tends to under-report induction, particularly

if the protein has not had several hours to mature prior to fixation. Thus, like all proxies for

gene expression, the results obtained using mCherryOpt must be interpreted with caution.

Plasmid stability and copy number

Our mCherryOpt vectors contain a pCD6 replicon. Although pCD6-based plasmids are

widely used in C. difficile, the published estimates of their segregational stability vary widely

(Cartman and Minton, 2010; Heap et al., 2009; Purdy et al., 2002), and we could not find any

published estimates of copy number. To assess plasmid stability, we passaged JIR8094

exconjugants containing pRPF185, pDSW1728, and pRAN738 in TY media without antibiotic

selection. After 46 generations, ∼70% of the cells in the culture could form a colony on TY

plates containing Thi at 10 μg/ml (Table 3.5). This corresponds to a rate of plasmid loss of <1%

per generation, assuming progeny grew at equal rates with or without the plasmids.

To determine the plasmid copy number, C. difficile cells containing either pRPF185 or

pRAN416 (a pRPF185 derivative with mldA) were grown in TY medium with thiamphenicol to an

OD600 of ∼0.6. Total plasmid and chromosomal DNA were extracted using phenol-chloroform,

and the relative amounts of various plasmid-borne and chromosomal genes were determined by

qPCR. Three primer sets were used to target three genes: rpoB encodes a subunit of RNA

polymerase and resides on the chromosome, cat confers resistance to thiamphenicol and

resides on the plasmids, and mldA codes for a C. difficile cell division protein and is found on the

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chromosome and one of the plasmids, pRAN416. The ratio of cat to rpoB ranged from 4 to 7,

while the ratio of mldA to rpoB was closer to 10 (Table 3.6). Thus, we estimate that the plasmids

are present at approximately 4 to 10 copies per chromosome during exponential growth in TY.

Discussion

GFP and related fluorescent proteins are powerful tools for studying gene expression

and protein localization in a wide variety of organisms, but the fact that these reporters require

O2 for chromophore maturation has hampered their use in strict anaerobes. Nevertheless, we

recently used cyan fluorescent protein (CFP) to demonstrate septal localization of three new cell

division proteins in C. difficile (Ransom et al., 2014). This application was made possible by our

discovery that CFP produced during anaerobic growth can acquire fluorescence even after cells

are fixed to preserve their architecture. We show here that a codon-optimized mCherry red

fluorescent protein is superior to CFP because it matures quickly and there is less interference

from the intrinsic fluorescence of C. difficile.

We also constructed and characterized two plasmids that we think will prove generally

useful to the C. difficile community (Figure 3.2). pDSW1728 is designed to be used as a reporter

of gene expression because unique restriction sites for NheI and SacI facilitate placing

mCherryOpt under the control of essentially any promoter-regulatory region of interest.

pRAN473 is designed for protein localization studies and has an in-frame MCS that can be used

for fusing mCherryOpt to the N-terminus of target proteins. The copy number of these plasmids

is in the range of 4 to 10 per genome, and they are stably maintained even in the absence of

antibiotic selection.

For studies of promoter activity, mCherryOpt has a couple of advantages that should

make it a useful complement to alternatives such as the gus reporter and RT-PCR. First,

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mCherryOpt is convenient because no manipulations are necessary beyond a 2-h (or less)

exposure of the culture to air. The fact that fixation can be omitted for mCherryOpt should

facilitate some experiments, such as high-throughput screening of small-molecule libraries to

identify potential new drugs. However, to observe native patterns of protein localization

fixation is essential because C. difficile dies and lyses after exposure to air. Second, when

mCherryOpt is visualized by microscopy it can be used to ask whether gene expression is

uniform across all cells in a population. Although this is often the case, there are well-

documented exceptions that would be difficult to study using gus or RT-PCR, such as bistable

regulatory switches and the spatial variation of gene expression in biofilms (Dubnau and Losick,

2006; Lequette and Greenberg, 2005; Maamar et al., 2007; Stewart and Franklin, 2008).

Despite the clear utility of mCherryOpt, investigators need to be mindful of its

limitations. We found that mCherryOpt underestimated PpdaV transcription, particularly when

cells were analyzed after 1 h of induction. Fluorescent proteins are generally not well suited for

monitoring the kinetics of changes in gene expression because they fold slowly and have long

half-lives [e.g., >24 h for wild-type GFP and 40 to 190 min for “unstable” GFP variants developed

to facilitate studies of transient gene expression in bacteria (Andersen et al., 1998)]. When

using fluorescent proteins to study protein localization, it should be remembered that

heterologous tags can perturb protein function, sometimes in ways that are difficult to

recognize (Margolin, 2012). In addition, the requirement for O2 precludes use of mCherryOpt

for studies of dynamic protein localization in live C. difficile cells, although this should not be a

problem in anaerobes that tolerate transient exposure to air (Hartman et al., 2011). Some

alternative fluorescent labels currently under development are smaller and do not require O2, so

they may overcome these limitations (Mukherjee and Schroeder, 2014). The limitations of

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mCherryOpt notwithstanding, our findings suggest it will prove to be a useful addition to the C.

difficile molecular biology toolbox.

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Table 3.1. Strains used in chapter three.

Strain Genotype and description Reference or Source

E. coli OmniMAX –

2 T1R F´ [proAB+ lacIq lacZΔM15 Tn10(TetR) Δ(ccdAB)] mcrA Δ(mrr-hsdRMS-mcrBC) Φ80lacZΔM15 Δ(lacZYA-argF) U169 endA1 recA1 supE44 thi-1 gyrA96 relA1 tonA panD.

Invitrogen

XL1-Blue endA1 gyrA96(nalR) thi-1 recA1 relA1 lac glnV44 [F' proAB+ lacIq Δ(lacZ)M15] hsdR17(rK

- mK+) Tn10(TetR)]

Aligent

HB101/pRK24 F− mcrB mrr hsdS20(rB

− mB

−) recA13 leuB6 ara-14 proA2

lacY1 galK2 xyl-5 mtl-1 rpsL20.

(Trieu-Cuot et al., 1987)

EC3272 XL1-Blue / pDSW1728 (Ptet::mCherryOpt cat) This study RAN473 Omnimax / pRAN473 (Ptet::mCherryOpt –MCS cat) This study

C. difficile

JIR8094 Spontaneous erythromycin-sensitive derivative of strain 630.

(O'Connor et al., 2006)

TCD20 csfV mutant (Ho et al., 2014)

RAN346 JIR8094 / pRAN334 (Ptet::cfpopt cat) (Ransom et al., 2014)

RAN347 JIR8094 / pRAN332 (Ptet::gfpmut2 cat) This study RAN429 JIR8094 / pRAN416 (Ptet::mldA cat) This study RAN433 JIR8094 / pRPF185 (Ptet::gusA cat) (Ransom et

al., 2014) RAN445 JIR8094 / pDSW1728 (Ptet::mCherryOpt cat) This study RAN538 JIR8094 / pRAN534 (Ptet::mCherryOpt –zapACd cat) This study RAN539 JIR8094 / pRAN535 (Ptet:: mCherryOpt –mldA cat) This study RAN829 JIR8094 / pRAN738 (PpdaV:: mCherryOpt cat) This study RAN836 TCD20 / pRAN738 (PpdaV::mCherryOpt cat) This study

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Table 3.2. Oligonucleotide primers used in chapter three.

Oligo Purpose

Sequence 5`-3`a RP157 Cloning gfpmut2 into pRPF185

GGGGAGCTCCTGCAGTAAAGGAGAAAATTTTATGAGTAAAGGAGAAGAACTTTTCACTGG

RP158 Cloning gfpmut2 into pRPF185

CCCGGATCCTTATTTGTATAGTTCATCCATGCCATGTG

RP176 Cloning zapA into pRAN357 AAAGCATGCATGAACAAAGTAATGGTTAAAATCCATGG

RP177 Cloning zapA into pRAN357 TTTGGCGCGCCTTATTCCACATTTTTTGCATCATTATTTAACC

RP204 Cloning mCherryOpt with MCSb into pRPF185

GGGGAGCTCCTGCAGTAAAGGAGAAAATTTTATGG

RP203 Cloning mCherryOpt with MCSb into pRPF185

GGCGGATCCGGCGCGCCTCAGCTGTTTAATTAAGTCGACGCATGCGTTCATTTTATATAATTCATCCATACCTCC

RP206 mCherryOpt sequencing primer CAAATTCATGTCCATTAACAGATCCTTCC RP207 mCherryOpt sequencing primer

GCATATAATGTTAATATTAAATTAGATATAAC

RP306 Cloning pdaV promoter in front of mCherryOpt

AAAAGCTAGCATGTGGCAAATAGTTGTTTTGCTATTTATTATTTG

RP307 Cloning pdaV promoter in front of mCherryOpt

TTTGAGCTCTACATTTATATTTTTGTAGTATTTTATCCCAAAAATTTACAC

TEQ009 qPCR primer for rpoB

AAGAGCTGGATTCGAAGTGCGTGA

TEQ010 qPCR primer for rpoB

ACCGATATTTGGTCCCTCTGGAGT

RP314 qPCR primer for catR GAAGGTTGACCACGGTATCAT

RP315 qPCR primer for catR CGCAACGGTATGGAAACAATC

RP316 qPCR primer for mldA AGTGGTTATTGTTGGTGTAGGA

RP317 qPCR primer for mldA

GCTTGTTGCTGAGTTGATGA

RP376 qPCR primer for pdaV TGTTCGCGTCAGCTCTTT

RP377 qPCR primer for pdaV ACTTGGCCCTTTACTAACTTCT

RP378 qPCR primer for mCherryOpt AGAAGGAGAAGGAGAAGGAAGA

RP379 qPCR primer for mCherryOpt AATGGTAATGGACCACCTTTAGT

99

a Restriction sites underlined.

b MCS, multicloning site.

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Table 3.3. Plasmids used in chapter three.

Plasmid Relevant Features Reference

pGFPmut2 gfpmut2 (Cormack et al.,

1996) pMA-T-mCherryOpt Synthesized mCherryOpt This study

pRPF185 E. coli-C. difficile shuttle vector with tetracycline-inducible promoter. Ptet::gusA

cat CD6ori RP4oriT-traJ pMB1ori

(Fagan and Fairweather,

2011)

pDSW1728 Ptet::mCherryOpt cat This study pRAN332 Ptet::gfpmut2 cat This study pRAN334 Ptet::cfpopt cat (Ransom et al.,

2014) pRAN357 Ptet:: cfpopt–MCS cat (Ransom et al.,

2014) pRAN409 Ptet:: cfpopt-zapACd This study pRAN410 Ptet:: cfpopt-mldA (Ransom et al.,

2014) pRAN416 Ptet::mldA (Ransom et al.,

2014) pRAN473 Ptet::mCherryOpt –MCS cat This study pRAN534 Ptet::mCherryOpt –ZapACd cat This study pRAN535 Ptet::mCherryOpt –MldA cat This study pRAN738 PpdaV::mCherryOpt cat This study

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Table 3.4. Codon usage in mCherryOpt.

Gene %GC Unfavorable codonsa No./total %

mCherry 62.4 215/236 91 mCherryOpt 30.1 3/236 1

mldA 26.8 134/847 16 zapA 31.6 49/196 25 rpoB 33.3 205/1238 17 tcdA 27.1 420/2710 15

a Codon preferences for C. difficile 630 were retrieved from http://www.kazusa.or.jp/codon/cgi-bin/showcodon.cgi?species=272563 (Nakamura et al., 2000). A codon was defined as unfavorable if it was at least 50% less common than the most frequently used codon for that amino acid in C. difficile (Sastalla et al., 2009). The GC-content of the C. difficile 630 genome is 29.1% (Sebaihia et al., 2006).

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Table 3.5. Plasmid stability.

Plasmid % ThiRa % Stability b

pRAN738 80.9 ± 1.6 99.5 ± 0.0 pDSW1728 78.9 ± 2.9 99.5 ± 0.1

pRPF185 66.8 ± 20.5 99.1 ± 0.7 a The fraction of cells resistant to thiamphenicol. Mean ± SD of three experiments.

b The fraction of daughter cells that retain the plasmid after each cycle of cell division, as

calculated based on ThiR data (see Materials and Methods). Mean ± SD of three experiments.

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Table 3.6. Plasmid copy number.

Ratioa Target genes RAN433b RAN429c catR / rpoB 4.5 ± 0.3 7.3 ± 0.9 mldA / rpoB 1.4 ± 0.9 9.6 ± 0.9 catR / mldA 4.0 ± 2.3 0.76 ± 0.1

a Mean ± SD of 3 biological replicates, each assayed with 3 technical replicates.

b RAN433 harbors pRPF185: cat is on the plasmid, rpoB and mldA are chromosomal.

c RAN429 harbors pRAN416: cat is on plasmid, rpoB is on the chromosome, and mldA is on both

the plasmid and the chromosome.

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M V S K G E E D N M A I I K

GAGCTCCTGC AGTAAAGGAG AAAATTTTAT GGTATCTAAA GGAGAAGAAG ATAATATGGC TATAATTAAA E F M R F K V H M E G S V N G H E F E I E G E G GAATTTATGA GATTTAAAGT TCATATGGAA GGATCTGTTA ATGGACATGA ATTTGAAATA GAAGGAGAAG E G R P Y E G T Q T A K L K V T K G G P L P F GAGAAGGAAG ACCTTATGAA GGTACTCAAA CAGCTAAATT AAAAGTAACT AAAGGTGGTC CATTACCATT A W D I L S P Q F M Y G S K A Y V K H P A D I TGCATGGGAT ATATTAAGTC CTCAATTTAT GTATGGATCT AAAGCATATG TTAAACATCC TGCTGATATA P D Y L K L S F P E G F K W E R V M N F E D G G CCTGATTATT TAAAATTATC ATTTCCAGAA GGTTTTAAAT GGGAAAGAGT TATGAATTTT GAAGATGGTG V V T V T Q D S S L Q D G D F I Y K V K L R G GAGTAGTTAC TGTAACACAA GATAGTTCAT TACAAGATGG AGATTTTATA TATAAAGTTA AATTAAGAGG T N F P S D G P V M Q K K T M G W E A S S E R TACTAATTTT CCATCTGATG GACCAGTAAT GCAAAAGAAA ACTATGGGAT GGGAAGCATC ATCTGAAAGA M Y P E D G A L K G E I K Q R L K L K D G G H Y ATGTATCCAG AAGATGGAGC ATTAAAAGGT GAAATTAAAC AAAGATTAAA ATTAAAAGAT GGAGGTCATT D A E V K T T Y K A K K P V Q L P G A Y N V N ATGATGCTGA AGTTAAGACT ACATATAAAG CTAAGAAACC AGTTCAATTA CCAGGTGCAT ATAATGTTAA I K L D I T S H N E D Y T I V E Q Y E R A E G TATTAAATTA GATATAACTT CTCATAATGA AGATTATACT ATAGTTGAAC AATATGAAAG AGCAGAAGGA R H S T G G M D E L Y K * AGACATTCAA CAGGAGGTAT GGATGAATTA TATAAATAAG GATCC

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Figure 3.1. DNA sequence of mCherryOpt. Shown is a synthetic 735 bp DNA fragment encoding

the mCherryOpt gene that was codon-optimized for expression in low GC bacteria. The

predicted amino acid sequence is shown in single-letter code above the DNA. Restriction sites

for SacI and BamHI are underlined and the Shine-Dalgarno sequence is double-underlined and in

bold face (Vellanoweth and Rabinowitz, 1992).

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107

Figure 3.2. Genetic maps of C. difficile mCherryOpt plasmids. (A) pDSW1728, for studies of gene

expression. Expression of mCherryOpt is under Ptet control, but the promoter-regulatory region

can be replaced using the NheI and SacI restriction sites. (B) pRAN473, for studies of protein

localization. (C) Multicloning site (MCS) in pRAN473. The last five amino acids derived from

mCherryOpt are shown in boldface above the DNA sequence. Both plasmids are derivatives of

pRPF185. Features depicted: mCherryOpt or mCherryOpt-MCS encodes mCherryOpt protein

codon optimized for expression in low-GC bacteria; tetR encodes the Tet repressor from Tn10;

orfB and repA, the replication region; ColE1 ori, replication region of the E. coli plasmid pBR322

modified for higher copy number; catP, the chloramphenicol acetyltransferase gene from

Clostridium perfringens, conferring resistance to thiamphenicol in C. difficile or chloramphenicol

in E. coli; traJ, encodes a conjugation transfer protein from plasmid RP4.

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109

Figure 3.3. Comparison of fluorescent proteins and autofluorescence. The top row of

micrographs shows C. difficile exconjugants producing mCherryOpt from pDSW1728 (A), CFPopt

from pRAN334 (B), or GFPmut2 from pRAN332 (C). Cells were photographed using filters

appropriate for each protein as indicated above the images. The middle row shows C. difficile

carrying pRPF185, which does not produce any fluorescent protein, photographed with the

same filter sets (D, red; E, blue; F, green). The bottom row shows phase-contrast micrographs

corresponding to images D to F.

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111

Figure 3.4. Time course of fluorescence development. (A) C. difficile exconjugants producing

mCherryOpt from pDSW1728 were induced with 400 ng of ATc per ml, fixed, removed from the

anaerobic chamber, and photographed at the times indicated. “Prior” refers to an unfixed

sample photographed immediately after removal from the anaerobic chamber. (B) C.

difficile/pDSW1728 (mCherryOpt) or C. difficile/pRPF185 (Neg) were grown and induced as

described above. Cells were fixed or not as indicated, transferred to PBS, and removed from the

anaerobic chamber. Fluorescence (red filter) was recorded using a plate reader at 15-min

intervals and normalized to the OD600 at time zero. The data points represent the means and

standard deviations of three independent cultures per group, all grown and processed on the

same day. These results are representative of three experiments.

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113

Figure 3.5. Septal localization of mCherryOpt fusions to the division proteins MldA and ZapA. C.

difficile cells harboring pRAN535 (mCherryOpt-MldA) or pRAN534 (mCherryOpt-ZapACd) were

fixed, removed from the anaerobic chamber, and photographed the next morning. (A and B)

Phase-contrast images. (A′ and B′) Fluorescence images. Arrows indicate examples of septal

localization. (C and D) Quantitation of fluorescence intensity along transects through the cells

marked with an asterisk in panels A′ and B′. The transects begin ∼1 μm before each cell (i.e.,

the end with the asterisk) and extend ∼1 μm beyond each cell. As noted elsewhere (Ransom et

al., 2014) and as documented in Figure 3.8 in the supplemental material, C. difficile does not

always divide precisely at the midcell.

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115

Figure 3.6. Multiple sequence alignment of ZapA orthologs. Alignment made with Clustal

Omega (Sievers et al., 2011) using default parameters and the following sequence accession

numbers: E. coli, NP_417386.1, B. subtilis, NP_390739, C. difficile YP_001087177.1. The E. coli

sequence was truncated at residue 91 (out of 109 total) and the C. difficile sequence was

truncated at residue 98 (out of 196 total).

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117

Figure 3.7. mCherryOpt-ZapACd localizes to division sites. To determine the position of division

sites, wild-type C. difficile was grown to mid-log phase in TY, removed from the anaerobic

chamber, stained with the membrane dye FM4-64 as described (Ransom et al., 2014), and

immediately photographed under fluorescence. ImagePro software was used to measure in

pixels the total cell length and the distance from the nearest pole to the septum. The distance

to the septum was then expressed as a fraction of cell length and graphed such that 0

represents the pole and 0.5 represents the midcell. For clarity, cells were binned into intervals

of 0.05 units. The positions of MldA and ZapACd were determined similarly using C. difficile cells

harboring pRAN535 (mCherryOpt-mldA) or pRAN534 (mCherryOpt-zapACd). These cells were

fixed, removed from the anaerobic chamber, and photographed the next morning under

fluorescence. In all cases at least 150 cells were scored. Cells without an obvious septum or

with multiple septa (chains of cells) were excluded from the analysis. The results show that (a)

most septa as determined using FM4-64 are very near the midcell and (b) the positional

distribution of mCherryOpt-MldA and mCherryOpt-ZapACd fluorescent bands is essentially

identical to that of septa.

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119

Figure 3.8. Use of mCherryOpt to monitor induction of the pdaV promoter by lysozyme. Wild-

type (A) or csfV mutant (B) strains of C. difficile containing pRAN738 (PpdaV::mCherryOpt) were

grown to an OD600 of 0.3, exposed to lysozyme for 1 h, fixed, and removed from the anaerobic

chamber. Fluorescence and phase contrast micrographs were obtained 16 h later. (C) Same as

in panels A and B, except that fluorescence was quantified using a plate reader and normalized

to the OD600.

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121

Figure 3.9. Induction of PpdaV::mCherryOpt as measured by fluorescence and qRT-PCR. Wild-

type C. difficile containing pRAN738 (PpdaV::mCherryOpt) was exposed to lysozyme for 1 h or 4 h.

Abundance of pdaV and mCherryOpt mRNA was assessed directly by qRT-PCR (with

normalization to rpoB). Abundance of mature mCherryOpt protein was determined using a

fluorescence reader (with normalization to OD600). Values are graphed as the mean and

standard deviation of three technical replicates from a single experiment and are representative

of two experiments. pdaV expression was only analyzed at the 1 h time point. Note that

fluorescence under-reports induction, especially at the earlier time point, suggesting

mCherryOpt folding/maturation is slow. Note also that pdaV and mCherryOpt mRNA were

induced similarly, which indicates the ~300 bp promoter fragment cloned onto the plasmid

carries all of the information needed for proper regulation of pdaV transcription under these

condition.

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CHAPTER IV

Toxin gene expression is bistable in Clostridium difficile

Introduction

Clostridium difficile is a Gram-positive, spore-forming, anaerobic bacterium and

opportunistic pathogen. C. difficile infections can cause antibiotic-associated diarrhea and

progress to life-threatening conditions, including pseudomembranous colitis and toxic

megacolon. Disease is mediated primarily through two exotoxins known as TcdA and TcdB

(Lyerly et al., 1982; Lyerly et al., 1985; Triadafilopoulos et al., 1987; Voth and Ballard, 2005).

Both are glucosyltransferases that glucosylate host proteins, particularly the Rho family of

GTPases (Carter et al., 2011a; Dingle et al., 2008; Just et al., 1994; Olling et al., 2011; Rupnik et

al., 2005; Sauerborn et al., 1997). This leads to collapse of the actin cytoskeleton and loss of

tight junctions (Feltis et al., 2000; Gerhard et al., 2008; Hartmann et al., 2003; Heine and Barth,

2006; Jank et al., 2007b; Just et al., 1995c; Nusrat et al., 1996; Warny et al., 1997). The end

result is gastrointestinal distress.

TcdA and TcdB are encoded on a 19.6 kb pathogenicity locus (PaLoc), along with three

additional toxin-related genes: tcdR, tcdC, and tcdE (Figure 4.1) (Cohen et al., 2000). TcdR is

absolutely required for tcdA and tcdB expression (Mani and Dupuy, 2001). TcdR is an alternative

sigma factor that recruits RNA polymerase to the promoters for tcdA and tcdB, and to its own

promoter. The roles of TcdC and TcdE are controversial. TcdC is proposed to function as an

anti-sigma for TcdR and thus negatively regulates toxin production (Carter et al., 2011b; Dupuy

et al., 2008; Matamouros et al., 2007), but this finding has been challenged (Cartman et al.,

2012; Murray et al., 2009). TcdE is a predicted membrane protein with some similarity to the

holins that create pores for bacteriophage to escape the cytoplasm (Carter et al., 2011b; Murray

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et al., 2009). But whereas two reports suggest TcdE is required for toxin secretion (Govind and

Dupuy, 2012; Tan et al., 2001), another study found TcdE to be completely dispensable (Olling et

al., 2012).

Toxin regulation is one of the best-studied aspects of C. difficile biology. When C.

difficile is grown in the laboratory in rich media, toxin is produced upon entry into stationary

phase (Karlsson et al., 1999), suggesting expression of tcdA and tcdB responds to nutrient

limitation. Consistent with this view, readily metabolized carbon sources like glucose and a

variety of amino acids also reduce toxin production (Dupuy and Sonenshein, 1998; Karlsson et

al., 1999; Karlsson et al., 2003; Karlsson et al., 2000). These effects are thought to be mediated

in part by the alternative sigma factor TcdR (also known as TcdD), whose expression is

influenced by nutrient availability and temperature (Karlsson et al., 2003).

Toxin gene expression in C. difficile is also influenced by several global regulators

(summarized in Figure 4.1). CodY is widely distributed in Firmicutes and functions as a repressor

when both GTP and branched chain amino acids are abundant in the cell (Guedon et al., 2001;

Ratnayake-Lecamwasam et al., 2001). In C. difficile, CodY represses 146 genes, including all five

genes in the PaLoc (Dineen et al., 2010). In vitro CodY binds to the promoters for tcdA, tcdB,

tcdC, and tcdR, but because affinity is about 10-folder higher for the tcdR promoter, this is likely

to be the most important target for CodY regulation of toxin production (Dineen et al., 2007).

CcpA, or carbon catabolite protein A, is a global regulator that responds to catabolizable

carbohydrates like glucose (Deutscher et al., 2008; Johnson et al., 2009). In C. difficile, CcpA

regulates ~140 genes (Antunes et al., 2012) and binds directly to the promoter regions of tcdA,

tcdB, tcdC, and tcdR (Antunes et al., 2011a). Because CcpA has ~10-fold higher affinity for the

tcdR promoter than for the other promoters, it is thought to work primarily through controlling

expression of TcdR (Antunes et al., 2011a). A number of other transcriptional regulators have

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been shown to impact toxin gene expression: PrdR (Bouillaut et al., 2013), SigD (El Meouche et

al., 2013; McKee et al., 2013), Agr (Martin et al., 2013), Spo0A (Mackin et al., 2013), SigH (Saujet

et al., 2011), and FliA (Aubry et al., 2012). These are less well-understood, and some seem to be

restricted to certain C. difficile strains.

Previous studies of toxin production have used bulk samples of cells or media, and thus

reflect the average behavior of the cells in the culture. Our development of mCherryOpt as a

reporter for gene expression in C. difficile (Chapter 3) enables analysis of toxin expression at the

level of individual cells. Here, we used mCherryOpt to study toxin regulation in C. difficile.

Remarkably, we found toxin gene expression is bistable; in stationary phase, the population

bifurcates into a group of cells that is “TcdA-ON” and a group that is “TcdA-OFF.” About 30% of

the cells are in the TcdA-ON state, and the mean fluorescence intensity of these cells is about

50-fold higher than the TcdA-OFF cells. Follow-up studies indicate expression of tcdR is the

genetic switch that determines whether or not a cell produces toxin.

Materials and Methods

Strains, media, and growth conditions

Bacterial strains used in this study are listed in Table 4.1. This study included five wild-

type C. difficile strains: JIR8094, R20291, CD196, NAP07, and NAP08. C. difficile mutants were

derived from the erythromycin sensitive isolate JIR8094, a derivative of the sequenced strain

CD630 (O'Connor et al., 2006; Sebaihia et al., 2006). C. difficile was routinely grown in Tryptone

Yeast (TY) media, supplemented as needed with thiamphenicol at 10 μg/ml, erythromycin at 5

μg/ml, kanamycin at 50 μg/ml, or cefoxitin at 16 μg/ml. TY consisted of 3% tryptone, 2% yeast

extract, and 2% agar (for solid media). TY included 0.1% L-cysteine during routine maintenance

of C. difficile cultures, but cysteine was generally omitted when assaying toxin production. C.

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difficile strains were maintained at 37°C in an anaerobic chamber (Coy Laboratory products) in

an atmosphere of 10% H2, 5% CO2, and 85% N2.

Escherichia coli strains were grown in LB medium at 37°C with chloramphenicol at 20

μg/ml and ampicillin at 100 μg/ml as needed. LB contained 10% tryptone, 5% yeast extract, 1%

NaCl and, for plates, 1.5% agar.

Plasmid and strain construction

All plasmids are listed in Table 4.2. Regions of plasmids constructed using PCR were

verified by DNA sequencing. The oligonucleotide primers used in this work were synthesized by

Integrated DNA Technologies (Coralville, IA). Primers are listed in Table 4.3.

The C. difficile null mutant of tcdR was constructed using modified TargeTron

procedures (Sigma-Aldrich) to insert a group II intron conferring Erm resistance (Heap et al.,

2010). Primers for retargeting the group II intron were designed using the ClosTron algorithm

(Heap et al., 2010). To retarget the intron to insert after nucleotide 142 of tcdR, the intron

template was amplified by PCR as outlined in the TargeTron user manual (Sigma-Aldrich) using

an EBS universal primer designated CDE914 in combination with primers RP398, RP399, and

RP400. The resulting PCR product and the vector pBL100 (Bouillaut et al., 2013) were digested

with HindIII and BsrGI, and then ligated to create plasmid pRAN1034. pRAN1034 was

transformed into the E. coli conjugation donor strain HB101/pRK24 (Trieu-Cuot et al., 1987).

Then pRAN1034 was transferred to C. difficile JIR8094 via conjugation and isolates in which the

intron had moved to the tcdR locus were obtained by selection for Erm-resistance as described

(Heap et al., 2010; Ho and Ellermeier, 2011; Ransom et al., 2015; Ransom et al., 2014). Intron

insertion into tcdR was confirmed by PCR using primer pair TEQ111/RP407. Finally, loss of the

TargeTron plasmid was confirmed by thiamphenicol sensitivity. The codY null mutant was built

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similarly. Primers RP323-RP325 were used to generate the plasmid pCE536. This mutant was

confirmed using primers RP401 and RP402.

For expression studies we constructed plasmids with promoters from tcdA, tcdB, and

tcdR. The plasmids are all derivatives of pDSW1728, which has a tetracycline-inducible

promoter and mCherryOpt (Ptet::mCherryOpt) (Ransom et al., 2015). Promoters were amplified

using the following primer sets: PtcdA (RP304 and RP305), PtcdB (RP345 and RP346), and PtcdR

(RP347 and RP348). The PCR products were digested with NheI and SacI, then ligated into

pDSW1728 digested with the same enzymes. The resulting plasmids were designated pRAN737

(PtcdA), pRAN841 (PtcdB), and pRAN842 (PtcdR). These plasmids were constructed using OmniMax-

2 T1R as the cloning host, transformed into HB101/pRK24, and then introduced into C. difficile

strains by conjugation.

To regulate tcdR expression in C. difficile, we built two constructs that had tcdR under an

inducible promoter: Ptet (Fagan and Fairweather, 2011) or PpdaV (Ransom et al., 2015). To build

PpdaV::tcdR, a synthetic DNA fragment (gBlock) containing both the promoter and gene was

synthesized by Integrated DNA Technologies, and the fragment was amplified using RP374 and

RP375. This DNA was digested with XmaI and inserted into the XmaI site of pRAN837. The

resulting plasmid was designated pRAN1018 (PtcdA::mCherryOpt; PpdaV::tcdR). To build Ptet::tcdR,

the Ptet promoter was amplified by PCR using primers RP393 and RP394 , with pRPF185 as the

template (Fagan and Fairweather, 2011). The Ptet promoter was then swapped with the PpdaV

promoter in pRAN1018 using a KpnI and SphI digest and ligation. The resulting plasmid was

named pRAN1032 (PtcdA::mCherryOpt; Ptet::tcdR).

Fixation protocol

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Cells were fixed as previously described (Ransom et al., 2015). Briefly, a 500-μl aliquot

of cells in growth medium was added directly to a microcentrifuge tube containing 120 μl of a

5X fixation cocktail: 100 μl of 16% (wt/vol) paraformaldehyde aqueous solution (methanol-free;

catalog no. AA433689M; Alfa Aesar, Ward Hill, MA) and 20 μl of 1 M NaPO4 buffer (pH 7.4). The

sample was mixed, allowed to sit for 15 min, removed from the Coy chamber, and incubated on

ice for 45 min. The fixed cells were washed three times with PBS, resuspended in 30 μl of PBS,

and left in the dark at 4°C to allow chromophore maturation.

Microscopy

Microscopy was performed as described previously (Ransom et al., 2015). Cells were

immobilized using thin agarose pads (1%). Phase-contrast and fluorescence micrographs were

recorded on an Olympus BX60 microscope equipped with a ×100 UPlanApo objective (numerical

aperture, 1.35). For mCherryOpt the filter set (catalog no. 41004) comprised a 538- to 582-nm

excitation filter, a 595-nm dichroic mirror (long pass), and a 582- to 682-nm emission filter. This

filter set was from Chroma Technology Corp. (Brattleboro, VT). The fluorescence micrographs

were captured using 3-s exposure times, unless indicated otherwise. To ensure comparability of

fluorescence micrographs, the display range option was adjusted identically for all images.

Micrographs were cropped, and figures were assembled in Adobe Illustrator (Adobe Systems,

Inc., San Jose, CA).

Flow cytometry

Cells were analyzed at the Flow Cytometry Facility at the University of Iowa. The

equipment used in this study includes the Becton Dickinson LSR II with a 561nm laser and the

Becton Dickinson Aria II.

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Plate reader

The plate reader was used as described previously (Ransom et al., 2015). Briefly,

fluorescence and absorbance (OD600) was measured with an Infinite M200 Pro plate reader

(Tecan, Research Triangle Park, NC). Samples were prepared by adding 20 μl of fixed cells in PBS

to 180 μl of PBS to the well of a flat-bottom 96-well microtiter plate (AS Plate-PS-96-F-C; AG

Advangene, IL). Fluorescence was recorded as follows: excitation, 554 nm; emission, 610 nm;

gain setting, 100. The cell density (OD600) was also recorded and used to normalize the

fluorescence reading.

RNA isolation from fixed cells

RNA was extracted from fixed cells that had been sorted based on fluorescence

intensity. The two distinct populations were separated by 1.8 x103 fluorescent units. We

collected ~107 cells for each population. To maximize RNA recovery, the cells were first

concentrated by centrifugation with a tabletop centrifuge at full speed. The cells were

resuspended in 200 μl of TE buffer plus 5 μl of Proteinase K (800 units/ml, New England Biolabs)

and transferred to a 0.5 ml microcentrifuge tube. A thermocycler was used to heat the sample

to 50°C for 15 min then 80°C for 15 min. The RNA was purified using the RNeasy mini kit

(Qiagen) and recommended DNAse-treatment step. RNA was eluted off the column using 30 µl

of water. The concentration and quality of RNA was determined using a Bioanalyzer 2100 in the

Genomics Facility at the University of Iowa.

qRT-PCR

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Quantitative real-time polymerase chain reaction (qRT-PCR) experiments were

performed as previously described (Ho and Ellermeier, 2011; Ransom et al., 2015) using Sybr

green master mix (Applied Biosystems) and the following gene-specific primer pairs: tcdA

(TEQ121-TEQ122) and cat (RP314-RP315). Data were normalized to mRNA levels of the C.

difficile housekeeping gene rpoB (primer pair TEQ009-TEQ010).

Results

tcdA expression is bistable in C. difficile

Toxin gene expression is known to increase during stationary phase (Dupuy and

Sonenshein, 1998; Ketley et al., 1984; Osgood et al., 1993). To visualize toxin gene expression in

single cells, we introduced a PtcdA::mCherryOpt reporter plasmid into R20291 ribotype 027. As

expected, in log phase overall fluorescence of the culture as measured with a plate reader was

low and fluorescence microscopy revealed the vast majority of the cells were dark (e.g., sample

#1 in Figure 4.2A, B, C, D). Upon entry into stationary phase overall fluorescence increased ~5

fold and microscopy revealed a striking mixture of bright and dark cells (e.g., sample #5 in Figure

4.2A, B, C, D). Flow cytometry confirmed that the distribution of fluorescence intensities was

bimodal, indicative of two distinctly different subpopulations, TcdA-ON and TcdA-OFF. The

TcdA-ON fraction reached a maximum of ~40% of the cells in sample #5 (9 hours after

subculturing, ~5 hours after entering stationary phase) and declined slowly thereafter.

Multiple lines of evidence rule out the potential artifact of plasmid segregation. First,

the plasmid is reported to be very stable (Heap et al., 2009; Ransom et al., 2015). Second, a

virtually identical plasmid with a lysozyme-induced PpdaV::mCherryOpt (Ransom et al., 2015)

insert provided uniform red fluorescence across the population after exposure of the cells to

lysozyme [See (+), for positive control in Figure 4.2B, C, D]. As a negative control, a plasmid

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lacking mCherryOpt (pRPF185) failed to produce any red fluorescent signal [See (-) in Figure

4.2B, C, and D]. Finally, and most compelling, we sorted paraformaldehyde-fixed cells by

fluorescence activated cell sorting based upon expression levels of PtcdA::mCherryOpt into two

populations: TcdA-ON or TcdA-Off (Figure 4.3). After reversal of the crosslinks, RNA was isolated

from these cells and cDNA was made. We then performed qRT-PCR using rpoB mRNA for

normalization. Expression of tcdA from the chromosome was ~15-fold higher in the TcdA-ON

population compared to the TcdA-OFF population (Figure 4.3). In contrast the expression of

catR, which is encoded on the plasmid, was the same in both populations (Figure 4.3), further

demonstrating that the observed differences in mCherryOpt levels are not a consequence of

plasmid loss.

Expression of tcdA is bistable in multiple C. difficile ribotypes

The tcdA promoter region is highly conserved across different C. difficile isolates (Figure

4.4). To ask whether bistable expression of tcdA is a general property conserved in other C.

difficile strains, we introduced the PtcdA::mCherryOpt reporter plasmid into four additional C.

difficile strains: JIR8094, which is an erythromycin sensitive derivative of CD630 (ribotype 012);

CD196 (ribotype 027); NAP07 (ribotype 078); and NAP08 (ribotype 078). The CD196, NAP07,

and NAP08 strains are representative clinical isolates corresponding to the most commonly

isolated ribotypes of C. difficile (Walker et al., 2013; Wilcox et al., 2012). Expression of the

PtcdA::mCherryOpt fusion was bistable in all strains (Figure 4.5), demonstrating bistability

pertains to a range of clinically relevant C. difficile ribotypes.

Glucose alters the bistable expression of tcdA

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Toxin production is influenced by the state of cellular metabolism [reviewed in

(Bouillaut et al., 2015)]. For example, exogenous glucose and cysteine reduce toxin production

during entry in stationary phase (Antunes et al., 2012; Karlsson et al., 2000). On the other hand,

exogenous butyric acid has been reported to increase toxin production (Karlsson et al., 1999). In

principle, different levels of toxin production could reflect changes in the fraction of cells that

are TcdA-ON, changes in the extent of induction in the TcdA-ON subpopulation, or some

combination of the two. We used our PtcdA::mCherryOpt reporter to examine the effect of

glucose, cysteine, and butyric acid on toxin production in cells grown in TY. As expected, glucose

and cysteine reduced toxin production, but we were unable to reproduce the previous report of

butyric acid-mediated activation despite testing a variety of concentrations (Figure 4.6 and data

not shown). Flow cytometry revealed that glucose, but not cysteine, reduced the fraction of

TcdA-ON cells (Figure 4.6). Thus, glucose and cysteine appear to repress toxin production by

different mechanisms.

Toxin bistability is dependent upon TcdR

Bistable patterns of gene expression can arise from cell-to-cell variation in the levels of a

transcriptional activator protein that is part of a positive feedback loop (Dubnau and Losick,

2006). Considering the various factors implicated in control of toxin production in C. difficile,

the alternative sigma TcdR is a promising candidate for the master regulator of bistability. TcdR

activates its own expression (Mani et al., 2002), so higher levels of TcdR will be self-reinforcing.

Moreover, basal expression of tcdR is very low (Mani and Dupuy, 2001), creating a situation in

which “noise” or stochastic cell-to-cell variation in TcdR levels can push a subset of cells over a

threshold that locks them into the ON state.

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We constructed a tcdR::erm null mutant by using TargeTron mutagenesis to insert a 1.7

kb group II intron containing an erythromycin resistance cassette (Figure 4.7). To analyze toxin

gene expression, we conjugated the PtcdA::mCherryOpt reporter plasmid into the tcdR::erm

background. There was no red fluorescence even in stationary phase (Figure 4.8A), confirming

previous reports that TcdR is essential for transcription of tcdA (Karlsson et al., 2003; Mani and

Dupuy, 2001; Mani et al., 2002). Next we introduced a Ptet::tcdR construct that allowed us to

regulate the level of TcdR in the cell by adding different amounts of the inducer

anhydrotetracycline (ATc) (Fagan and Fairweather, 2011). As expected, overall fluorescence and

the fraction of cells in the TcdA-ON state increased with increasing amounts of ATc, approaching

90% TcdA-ON when ATc was at 100 ng/ml (Figure 4.8B, C). Nevertheless, bistability was not

observed at any ATc concentration; rather, fluorescence increased uniformly with increasing ATc

(Figure 4.8C). Thus, breaking the auto-catalytic feedback loop involved in tcdR expression

prevents bistability, consistent with the hypothesis that cell-to-cell variation in TcdR levels is the

underlying cause of bistable TcdA production in a wild-type background.

Toxin bistability is not dependent upon CodY

The global repressor CodY is known to regulate expression of tcdR, tcdA, and tcdB

(Dineen et al., 2010; Dineen et al., 2007). We introduced the PtcdA::mCherryOpt reporter plasmid

in a codY::erm null mutant constructed using TargeTron methods (Figure 4.7). Inactivating codY

increased fluorescence about 30-fold (Figure 4.9A). This effect compares favorably with a

previous study showing inactivating codY increases tcdA mRNA about 50-fold (Dineen et al.,

2010). The absence of CodY does not abolish bistability, but does increase the ratio of TcdA-ON

to TcdA-OFF cells about 3-fold (Figure 4.9B). The more striking result is that in the TcdA-ON

population, the levels of mCherryOpt are increased well beyond those observed in wild-type

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TcdA-ON cells. In summary, CodY affects both the extent of tcdA induction and the fraction of

cells that become TcdA-ON, but it is not required for bistability per se.

Preliminary evidence for bistable expression of tcdB

Because tcdA expression is bistable and TcdR levels appear to be critical for establishing

bistability, we hypothesized that expression of the other toxin (TcdB) and expression of the

master regulator (TcdR) would be bistable. Unfortunately, cells carrying a PtcdB::mCherryOpt

reporter plasmid were not fluorescent (data not shown), likely because expression was below

our detection limit. The expression of tcdB is reported to be 10 to 100 fold lower than tcdA

(Martin et al., 2013; Merrigan et al., 2010). However, in a codY::erm mutant background,

expression of the PtcdB::mCherryOpt reporter was readily detected and bistable (Figure 4.9). We

were unable to assess whether production of TcdR is bistable because we could not detect any

fluorescence from a PtcdR::mCherryOpt reporter plasmid, even in a codY::erm mutant background

(Figure 4.9 and data not shown). Similar reporter constructs incorporating different amounts of

DNA from the tcdR promoter region were also non-fluorescent (not shown). This finding is not

too surprising because expression of tcdR is known to be lower than that of tcdA or tcdB (Dupuy

and Sonenshein, 1998).

Discussion

In C. difficile, pathogenesis is mediated primarily by two large exotoxins encoded in the

PaLoc, TcdA and TcdB. There has been a lot of effort expended to understand how production

of these toxins is controlled. Early studies found that the toxins are produced upon entry into

stationary phase (Dupuy and Sonenshein, 1998; Moncrief et al., 1997). This response is

mediated by a dedicated sigma factor (TcdR) and by a host of global regulatory proteins, most of

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which sense various aspects of metabolism [reviewed in (Bouillaut et al., 2015; Dineen et al.,

2010; Mani et al., 2002; Voth and Ballard, 2005)]. All of these studies have relied on methods

that reflect that average behavior of the cells in the population, under the (unstated)

assumption that toxin production is relatively uniform across the population. Here we have

used a fluorescent protein reporter, mCherryOpt, to visualize expression of tcdA in individual

cells. Our results indicate that during entry into stationary phase only a subset of C. difficile cells

produce TcdA. Preliminary results for TcdB indicate that it too is produced in a bistable manner.

One question raised by these findings is, how is bistability controlled? In other words,

what is the critical factor that determines whether a particular cell will produce toxin or not?

Previous studies of bistability in other bacteria have revealed two characteristics that make a

regulatory protein well-suited for controlling a bistable switch (Dubnau and Losick, 2006). One

is low-level basal expression so that stochastic variation can lead to excursions that tip the

balance between an ON and an OFF state. The other is a positive feedback loop that reinforces

transient increases in cellular abundance of the activator. In the case of C. difficile, the

alternative sigma TcdR fulfills both criteria (Dupuy and Sonenshein, 1998; Mani and Dupuy,

2001; Mani et al., 2002). In support of this notion, we found that graded expression of tcdR

using a tetracycline-inducible promoter prevents development of bistability. Instead, as more

inducer is added to the culture, toxin production increases uniformly across the cells in the

population. These findings imply that in a wild-type background tcdR expression is itself

bistable. Unfortunately, efforts to test this idea using an mCherryOpt reporter were not

successful, presumably owing to the very low level of tcdR expression, which was below our

detection limit. In the future it would be interesting to assay tcdR mRNA levels by qRT-PCR in

TcdA-ON and TcdA-OFF populations. These cell types can be isolated by sorting a strain that

carries a PtcdA::mCherryOpt reporter plasmid.

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A plethora of global regulators have been implicated in control of toxin production in C.

difficile (Antunes et al., 2011a; Dineen et al., 2010; Dineen et al., 2007; Dupuy and Sonenshein,

1998; El Meouche et al., 2013; Mackin et al., 2013; Saujet et al., 2011). Here we tested one of

these, CodY, which is known to repress several genes on the PaLoc. In the absence of CodY, the

fraction of cells producing TcdA increased modestly (~3x) while the level of toxin production in

those cells increased more dramatically (~10x). Thus, CodY is not required for bistability.

Another interesting question is, what is the benefit of bistability in the case of C. difficile

toxin production? It remains unclear how TcdA and TcdB are transported across the cell

envelope. One possibility is that cell lysis may be required for release of the toxins. If “suicide”

is needed for toxin delivery, presumably only a subset of cells would lyse and release toxin while

a second subset would reap the benefits. However, another potential selective pressure relates

to the need for C. difficile to form O2-tolerant spores to spread from one host to the next.

Sporulation and toxin production may be mutually (or temporally) exclusive processes in C.

difficile (Bouillaut et al., 2015; Saujet et al., 2011). In this case, one can imagine that a subset of

cells provoke diarrhea by producing toxin, while another subset develops into spores to

facilitate survival until a new host is found.

We hope a thorough understanding of toxin regulation may lead to improved

therapeutic treatments. In theory, one may be able to target only toxin-ON cells and thus drive

the evolution of C. difficile to a non-toxin producing, non-pathogenic commensal microbe.

Alternatively, being able to prevent toxin production is expected to relieve (or prevent)

symptoms of C. difficile disease without disrupting the microbiome.

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Acknowledgments

"The data presented herein were obtained at the Flow Cytometry Facility, which is a

Carver College of Medicine / Holden Comprehensive Cancer Center core research facility at the

University of Iowa. The Facility is funded through user fees and the generous financial support

of the Carver College of Medicine, Holden Comprehensive Cancer Center, and Iowa City

Veteran's Administration Medical Center." “Research reported in this publication was

supported by the National Center for Research Resources of the National Institutes of Health

under Award Number 1S10 RR027219.”

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Table 4.1. Strains used in chapter four.

Strain Genotype and description Reference or Source

E. coli OmniMAX –

2 T1R F´ [proAB+ lacIq lacZΔM15 Tn10(TetR) Δ(ccdAB)] mcrA Δ(mrr-hsdRMS-mcrBC) Φ80lacZΔM15 Δ(lacZYA-argF) U169 endA1 recA1 supE44 thi-1 gyrA96 relA1 tonA panD.

Invitrogen

XL1-Blue endA1 gyrA96(nalR) thi-1 recA1 relA1 lac glnV44 [F' proAB+ lacIq Δ(lacZ)M15] hsdR17(rK

- mK+) Tn10(TetR)]

Aligent

HB101/pRK24 F− mcrB mrr hsdS20(rB

− mB

−) recA13 leuB6 ara-14 proA2

lacY1 galK2 xyl-5 mtl-1 rpsL20.

(Trieu-Cuot et al., 1987)

EC3272 XL1-Blue / pDSW1728 (Ptet::mCherryOpt cat) (Ransom et al., 2015)

RAN473 Omnimax / pRAN473 (Ptet::mCherryOpt –MCS cat) (Ransom et al., 2015)

C. difficile JIR8094 Spontaneous erythromycin-sensitive derivative of strain

630. (Ribotype 012) (O'Connor et

al., 2006) CD196 Wild-type C. difficile strain from France. AKA (ribotype

027/BI/NAP1).

NAP07 Wild-type C. difficile strain NAP08 Wild-type C. difficile strain R20291 Wild-type C. difficile strain from UK outbreak. AKA

(ribotype 027/BI/NAP1).

RAN820 NAP08 / pRAN737 (PtcdA:: mCherryOpt cat) This study RAN828 JIR8094 / pRAN737 (PtcdA:: mCherryOpt cat) This study RAN829 JIR8094 / pRAN737 (PpdaV:: mCherryOpt cat) (Ransom et

al., 2015) RAN912 JIR8094 / pRAN841 (PtcdB:: mCherryOpt cat) This study RAN913 JIR8094 / pRAN842 (PtcdR:: mCherryOpt cat) This study RAN925 R20291 / pRAN737 (PtcdA:: mCherryOpt cat) This study RAN934 CD196 / pRAN737 (PtcdA:: mCherryOpt cat) This study

RAN1101 NAP07 / pRAN737 (PtcdA:: mCherryOpt cat) This study RAN1116 JIR8094 codY330::ltrB::ermB This study RAN1123 JIR8094 tcdR142::ltrB::ermB This study RAN1124 RAN1116 (codY mutant) / pRAN737 (PtcdA:: mCherryOpt

cat) This study

RAN1127 RAN1123 (tcdR mutant) / pRAN737 (PtcdA:: mCherryOpt

cat) This study

TCD20 JIR8094 csfV63::ltrB::ermB (Ho et al., 2014)

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Table 4.2. Plasmids used in chapter four.

Plasmid Relevant Features Reference

pRPF185 E. coli-C. difficile shuttle vector with tetracycline-inducible promoter. Ptet::gusA cat CD6ori RP4oriT-traJ pMB1ori

(Fagan and Fairweather,

2011) pBL100 E. coli-C. difficile shuttle vector for creating C.

difficile mutants using TargeTron mutagenesis. ltrB::ermB::RAM ltrA cat bla CD6ori RP4oriT pMBlori

(Bouillaut et al., 2013)

pDSW1728 Ptet::mCherryOpt cat (Ransom et al., 2015)

pCE536 pBL100 / codY330 This study pRAN737 pDSW1728 derivative with PtcdA::mCherryOpt This study

pRAN841 pDSW1728 derivative with PtcdB::mCherryOpt This study

pRAN842 pDSW1728 derivative with PtcdR::mCherryOpt This study

pRAN1018 PtcdA::mCherryOpt / PpdaV::tcdR This study

pRAN1032 PtcdA::mCherryOpt / Ptet::tcdR This study pRAN1034 pBL100 / tcdR142 This study

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Table 4.3. Oligonucleotide primers used in chapter four.

Oligo Purpose Sequence 5`-3`a CDE914 EBS universal primer CGAAATTAGAAACTTGCGTTCAGTAAAC DW2002 Sequencing secondary cloning site in

pRAN1018 TGAGTGAGCTGATACCGCTCG

DW2003 Sequencing secondary cloning site in pRAN1018

CAACTTGCCCACTTCGACTGC

NF1438 Reverse primer for sequencing inserts in pRPF185

GATCCAGCACACTGGCATCTTTTTATTTAGGGATTTCTCAC

RP206 mCherryOpt sequencing primer CAAATTCATGTCCATTAACAGATCCTTCC RP207 mCherryOpt sequencing primer

GCATATAATGTTAATATTAAATTAGATATAAC

RP298 Forward primer for sequencing inserts in pRPF185

CCCAAATCCTTACATCTCCCC

RP304 tcdA promoter AAAAGCTAGCCGTGATGAAGGACAAAATGATATAGAAAATAAG

RP305 tcdA promoter TTTGAGCTCGTATTATTATTTTTGATAATAAATCCACTTCTATAACC

RP314 qRT-PCR for cat GAAGGTTGACCACGGTATCAT RP315 qRT-PCR for cat CGCAACGGTATGGAAACAATC RP323 IBS for constructing codY330::erm

AAAAAAGCTTATAATTATCCTTAACTACCGTAGTAGTGCGCCCAGATAGGGTG

RP324 EBS1 for constructing codY330::erm

CAGATTGTACAAATGTGGTGATAACAGATAAGTCGTAGTATATAACTTACCTTTCTTTGT

RP325 EBS2 for constructing codY330::erm TGAACGCAAGTTTCTAATTTCGATTGTAGTTCGATAGAGGAAAGTGTCT

RP345 tcdB promoter AAAAGCTAGCCTTGTAATTAATGAGCTTAAAGAAATATTTACAATAG

RP346 tcdB promoter TTTGAGCTCTACATCTAAATGCTAAAACTCTTTTATATATCCTCCTTTC

RP374 Amplify PpdaV::tcdR from gBlock CCCCCGGGGCGGTACCCTAGAATTAATTG

RP375 Amplify PpdaV::tcdR from gBlock GCCCCGGGCGCGCCTTACAAGTTAAAATAATTTTC

RP378 qRT-PCR for mCherryOpt AGAAGGAGAAGGAGAAGGAAGA RP379 qRT-PCR for mCherryOpt AATGGTAATGGACCACCTTTAGT RP393 Amplify Ptet from pRPF185 TATGGTACCTTAAGACCCACTTTCAC RP394 Amplify Ptet from pRPF185

TTTGCATGCAGATCTGTTAACGCTACGATCAAGC

RP398 IBS for constructing tcdR142::erm AAAAAAGCTTATAATTATCCTTAGAAACCGATTTAGTGCGCCCAGATAGGGTG

RP399 EBS1 for constructing tcdR142::erm

CAGATTGTACAAATGTGGTGATAACAGATAAGTCGATTTAATTAACTTACCTTTCTTTGT

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RP400 EBS2 for constructing tcdR142::erm TGAACGCAAGTTTCTAATTTCGATTGTTTCTCGATAGAGGAAAGTGTCT

RP401 Confirm codY mutant GACAGTTCAGTTATAGAAGATGAG RP402 Confirm codY mutant CTTCTTCTAATTCTTCACCTATTGCTC RP407 Confirm tcdR mutant CTTCCTCAAAAACAGACTTACTTTG TEQ009 qRT-PCR for rpoB AAGAGCTGGATTCGAAGTGCGTGA TEQ010 qRT-PCR for rpoB ACCGATATTTGGTCCCTCTGGAGT TEQ111 qRT-PCR for tcdR & confirm tcdR mutant AGCAAGAAATAACTCAGTAGATGATT TEQ112 qRT-PCR for tcdR TTATTAAATCTGTTTCTCCCTCTTCA TEQ121 qRT-PCR for tcdA GGAGAAGTCAGTGATATTGCTCTTG TEQ122 qRT-PCR for tcdA

CAGTGGTAGAAGATTCAACTATAGCC

a Restriction sites underlined.

141

142

Figure 4.1. Schematic diagram of toxin regulation. Shown is the PaLoc, which encodes for the

two main toxins (TcdA and TcdB), the master regulator of toxin (TcdR), a putative holin for toxin

release (TcdE), and the putative anti-sigma factor (TcdC). Also depicted are several known

regulators of toxin production. CodY and CcpA are repressors that bind directly to the tcdR,

tcdA, tcdB, and tcdC promoters. Spo0A and SigH also repress toxin production but it is currently

unclear if the repression is direct. The alternative sigma factor SigD contributes to transcription

of tcdR and thus increases toxin production.

143

144

Figure 4.2. Bistable expression of PtcdA::mCherryOpt in C. difficile R20291. (A) Growth curve of

PtcdA::mCherryOpt strain. At the times indicated by the arrows, samples of the culture were fixed

with paraformaldehyde and exposed to air to allow red fluorescence to develop. (B) Specific

fluorescence of PtcdA::mCherryOpt population as determined using a plate reader. Numbers on

the X-axis refer to samples fixed at the time points indicated in (A). Also shown is a negative

control strain containing a Ptet::gus reporter plasmid (-) and a positive control strain carrying a

PpdaV::mCherryOpt reporter plasmid and induced with lysozyme for 30 min (+). The “P”

designation refers to a blank consisting of PBS with no bacteria. (C) Expression of

PtcdA::mCherryOpt as assessed by microscopy. Micrographs are paired: fluorescence (above) and

phase micrographs (below). (D) Expression of PtcdA::mCherryOpt as assessed by flow cytometry.

The x-axis label is mCherryOpt fluorescence in arbitrary units.

145

146

Figure 4.3. Expression of chromosomal tcdA is induced in TcdA-ON cells. Left panel: cells

collected as TcdA-ON or TcdA-OFF. Right panel: qRT-PCR analysis of sorted cells using primers

for catR, tcdA, and rpoB (for normalization). FSC-A refers to forward scatter.

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Bl1 AGTAATGGTAGATATAATAAAAATATTAACAAATAAAAAGTGTTATCCAAATAAGAATAG 60 CD196 AGTAATGGTAGATATAATAAAAATATTAACAAATAAAAAGTGTTATCCAAATAAGAATAG 60 R20291 AGTAATGGTAGATATAATAAAAATATTAACAAATAAAAAGTGTTATCCAAATAAGAATAG 60 CD630 AGTAATGGTAGATATAATAAAGATATTAACAAATAAAAAGTGTTATCCAAATAAGAATAG 60 NAP07 AGTACTGGTAGATATAATAAAGATATTAACAAATAAAAAGTGTTATCCAAATAAGAATAG 60 NAP08 AGTACTGGTAGATATAATAAAGATATTAACAAATAAAAAGTGTTATCCAAATAAGAATAG 60 ****.****************.************************************** Bl1 CTGAAAGTTATCATAATTCATGAAACTGATAATGAAAACGAGGGAGCAGATGCCAAGAGA 120 CD196 CTGAAAGTTATCATAATTCATGAAACTGATAATGAAAACGAGGGAGCAGATGCCAAGAGA 120 R20291 CTGAAAGTTATCATAATTCATGAAACTGATAATGAAAACGAGGGAGCAGATGCCAAGAGA 120 CD630 CTGAAAGTTATCATAATTCATGAAACTAATAATGAAAACGAGGGAGCAGATGCCAAGAGA 120 NAP07 CTGAAAGTTATCATAATTCATGAAACTGATAATGAAAACGAGGGAGCAGATGCCAAGAGA 120 NAP08 CTGAAAGTTATCATAATTCATGAAACTGATAATGAAAACGAGGGAGCAGATGCCAAGAGA 120 ***************************.******************************** Bl1 CACACAAGTATTAAATACATATAATTTCGAAGCAAGTGTTCATTACTATATGGATGACAA 180 CD196 CACACAAGTATTAAATACATATAATTTCGAAGCAAGTGTTCATTACTATATGGATGACAA 180 R20291 CACACAAGTATTAAATACATATAATTTCGAAGCAAGTGTTCATTACTATATGGATGACAA 180 CD630 CACACAAGTATTAAATACATATAATTTCGAAGCAAGTGTTCATTACTATATAGATGACAA 180 NAP07 CACACAAGTATTAAATACATATAATTTCGAAGCAAGTGTTCATTACTATATGGATGACAA 180 NAP08 CACACAAGTATTAAATACATATAATTTCGAAGCAAGTGTTCATTACTATATGGATGACAA 180 ***************************************************.******** Bl1 GGTAGTATATCAAACATTGGTTCACAAAGATGGTGCATGGTCAGTTGGTAAAATCTATTA 240 CD196 GGTAGTATATCAAACATTGGTTCACAAAGATGGTGCATGGTCAGTTGGTAAAATCTATTA 240 R20291 GGTAGTATATCAAACATTGGTTCACAAAGATGGTGCATGGTCAGTTGGTAAAATCTATTA 240 CD630 GGTAGTATATCAAACATTGGTTCACAAAGATGGTGCATGGTCAGTTGGTAAAATCTATTA 240 NAP07 GGTAGTATATCAAACATTGGTTCACAAAGATGGTGCATGGTCAGTTGGTAAAATCTATTA 240 NAP08 GGTAGTATATCAAACATTGGTTCACAAAGATGGTGCATGGTCAGTTGGTAAAATCTATTA 240 ************************************************************ Bl1 AGCTACATTAGTTACAGATATCACAAACTATAATAGTTAAACATAGAAATATGTGTAAAT 300 CD196 AGCTACATTAGTTACAGATATCACAAACTATAATAGTTAAACATAGAAATATGTGTAAAT 300 R20291 AGCTACATTAGTTACAGATATCACAAACTATAATAGTTAAACATAGAAATATGTGTAAAT 300 CD630 AGCTACATTAGTTACAGATATCACAAACTATAATAGTTAAACATAGAAATATGTGTAAAT 300 NAP07 AGCTACATTAGTTACAGATATCACAAACTATAATAGTTAAACATAGAAATATGTGTAAAT 300 NAP08 AGCTACATTAGTTACAGATATCACAAACTATAATAGTTAAACATAGAAATATGTGTAAAT 300 ************************************************************ Bl1 TGTGATGGAAATTATTCAAAAACACAAAAATACGTGATGAAGGACAAAATGATAGAGAAA 360 CD196 TGTGATGGAAATTATTCAAAAACACAAAAATACGTGATGAAGGACAAAATGATAGAGAAA 360 R20291 TGTGATGGAAATTATTCAAAAACACAAAAATACGTGATGAAGGACAAAATGATAGAGAAA 360 CD630 TGTGATGGAAATTATTCAAAAACACAAAAATACGTGATGAAGGACAAAATGATATAGAAA 360 NAP07 TGTGATGGAAATTATTCAAAAACACAAAAATACGTGATGAAGGACAAAATGATAGAGAAA 360 NAP08 TGTGATGGAAATTATTCAAAAACACAAAAATACGTGATGAAGGACAAAATGATAGAGAAA 360 ******************************************************.***** Bl1 ATAAGTATCAAACCTTAATAAATGATTTAATTGATAGTTTAAAAGTTATAGGAAAAATAT 420 CD196 ATAAGTATCAAACCTTAATAAATGATTTAATTGATAGTTTAAAAGTTATAGGAAAAATAT 420 R20291 ATAAGTATCAAACCTTAATAAATGATTTAATTGATAGTTTAAAAGTTATAGGAAAAATAT 420 CD630 ATAAGTATCAAACCTTAATAAATGATTTAATTGATAGTTTAAAAGTTATAGGAAAAATAT 420 NAP07 ATAAGTATCAAACCTTAATAAATGATTTAATTGATAGTTTAAAAGTTATAGGAAAAATAT 420 NAP08 ATAAGTATCAAACCTTAATAAATGATTTAATTGATAGTTTAAAAGTTATAGGAAAAATAT 420 ************************************************************ Bl1 ATAAAGAAATAAAAACATTAAAAAAATATAAGATATGTTTACAAATTACTATCAGACAAT 480 CD196 ATAAAGAAATAAAAACATTAAAAAAATATAAGATATGTTTACAAATTACTATCAGACAAT 480 R20291 ATAAAGAAATAAAAACATTAAAAAAATATAAGATATGTTTACAAATTACTATCAGACAAT 480 CD630 ATAAAGAAATAAAAACATTAAAAAAATATAAGATATGTTTACAAATTACTATCAGACAAT 480 NAP07 ATAAAGAAATAAAAACATTAAAAAAATATAAGATATGTTTACAAATTACTATCAGACAAT 480 NAP08 ATAAAGAAATAAAAACATTAAAAAAATATAAGATATGTTTACAAATTACTATCAGACAAT 480 ************************************************************

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Bl1 CTCCTTATCTAATAGAAGAGTCAATTAACTAATTGAGTATCTTTAAATTGAAATGTTAGG 540 CD196 CTCCTTATCTAATAGAAGAGTCAATTAACTAATTGAGTATCTTTAAATTGAAATGTTAGG 540 R20291 CTCCTTATCTAATAGAAGAGTCAATTAACTAATTGAGTATCTTTAAATTGAAATGTTAGG 540 CD630 CTCCTTATCTAATAGAAGAGTCAATTAACTAATTGAGTATCTTTAAATTGAAATGTTAGG 540 NAP07 CTCCTTATCTAATAGAAGAGTCAATTAACTAATTGAGTATCTTTAAATTGAAATGTTAGG 540 NAP08 CTCCTTATCTAATAGAAGAGTCAATTAACTAATTGAGTATCTTTAAATTGAAATGTTAGG 540 ************************************************************ Bl1 AAGTGATTTAAATATGAAAACTTAAATTATAAAAAATCAATATTAATTTATCTTTAAAAA 600 CD196 AAGTGATTTAAATATGAAAACTTAAATTATAAAAAATCAATATTAATTTATCTTTAAAAA 600 R20291 AAGTGATTTAAATATGAAAACTTAAATTATAAAAAATCAATATTAATTTATCTTTAAAAA 600 CD630 AAGTGATTTAAATATGAAAACTTAAATTATAAAAAATCAATATTAATTTATTTTTAAAAA 600 NAP07 AAGTGATTTAAATATGAAAACTTAAATTATAAAAAATCAATATTAATTTATTTTTAAAAA 600 NAP08 AAGTGATTTAAATATGAAAACTTAAATTATAAAAAATCAATATTAATTTATTTTTAAAAA 600 *************************************************** ******** Bl1 ATAGAAAGGAGTGTATAAGATTTATTTTCAAAGTTTAAAAACAAGAAAATCAATTTAAAT 660 CD196 ATAGAAAGGAGTGTATAAGATTTATTTTCAAAGTTTAAAAACAAGAAAATCAATTTAAAT 660 R20291 ATAGAAAGGAGTGTATAAGATTTATTTTCAAAGTTTAAAAACAAGAAAATCAATTTAAAT 660 CD630 ATAGAAAGGAGTGTATAAGATTTATTTTCAAAGTTTAAAAACAAGAAAATCAATTTAAAT 660 NAP07 ATAGAAAGGAGTGTATAAGATTTATTTTCAAAGTTTAAAAAAAAGAAAATCAATTTAAAT 660 NAP08 ATAGAAAGGAGTGTATAAGATTTATTTTCAAAGTTTAAAAAAAAGAAAATCAATTTAAAT 660 *****************************************.****************** Bl1 TTCAGAAGGAATAAATGTGGTTATAGAAGTGGATTTATTATCAAAAATAATAGTACTAGG 720 CD196 TTCAGAAGGAATAAATGTGGTTATAGAAGTGGATTTATTATCAAAAATAATAGTACTAGG 720 R20291 TTCAGAAGGAATAAATGTGGTTATAGAAGTGGATTTATTATCAAAAATAATAGTACTAGG 720 CD630 TTCAGAAGGAATAAATGTGGTTATAGAAGTGGATTTATTATCAAAAATAATAATACTAGG 720 NAP07 TTCAGAAGGAATAAATGTGGTTATAGAAGTGGATTTATTATCAAAAATAATAATACTAGG 720 NAP08 TTCAGAAGGAATAAATGTGGTTATAGAAGTGGATTTATTATCAAAAATAATAATACTAGG 720 ****************************************************.******* Bl1 AGGTTTTT 728 CD196 AGGTTTTT 728 R20291 AGGTTTTT 728 CD630 AGGTTTTT 728 NAP07 AGGTTTTT 728 NAP08 AGGTTTTT 728 ********

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Figure 4.4. Alignment of tcdA promoters from different C. difficile strains. The upstream region

of tcdA from different strains was aligned using Clustal Omega (Sievers et al., 2011). The C.

difficile strains are Bl1, CD196, R20291, CD630, NAP07, and NAP08.

150

151

Figure 4.5. Expression of PtcdA::mCherryOpt is bistable in multiple C. difficile strains. The

indicated strains of C. difficile harboring the PtcdA::mCherryOpt reporter plasmid were fixed after

20 hours of growth (corresponding to time point 7 in Figure 4.2A). The strains shown represent

the following ribotypes: JIR8094 (ribotype 012), CD196 (ribotype 027), NAP07 (ribotype 078),

and NAP08 (ribotype 078).

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A

153

Figure 4.6. Effect of glucose, cysteine, and butyric acid on TcdA production. The R20291 strain

containing PtcdA::mCherryOpt was grown in TY amended with 0.5% glucose, 0.1% cysteine, or 30

mM butyrate. (A) Growth was monitored by OD600. At the points indicated, samples were fixed

and specific fluorescence (which is total fluorescence normalized to OD) was determined using a

plate reader. (B) Cultures in stationary phase (20 h after inoculation) were fixed and analyzed

flow cytometry.

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155

Figure 4.7. PCR confirmation of C. difficile regulatory mutants. Genomic DNAs from the strains

indicated were used for PCR with oligonucleotide primers that anneal to regions flanking the

predicted insertion site. If the gene is intact a product of ~300 bp should be amplified. If the

erm resistance cassette is inserted into the gene, a larger product of about 2 kb is expected.

156

157

Figure 4.8. TcdR mediates bistable expression of tcdA. (A) TcdR is required for tcdA expression.

Wild-type/PtcdA::mCherryOpt and tcdR::erm/PtcdA::mCherryOpt were grown to stationary phase

(24 h). Samples of each culture were fixed and analyzed for red fluorescence by flow cytometry.

(B) Breaking the positive feedback loop that controls tcdR expression prevents development of

bistability. The tcdR::erm mutant harboring a plasmid with both Ptet::tcdR and PtcdA::mCherryOpt

was grown to mid-log phase (OD600 = 0.3), at which time ATc was added as indicated to induce

expression of tcdR. After 1 h, samples were fixed an analyzed using a plate reader (B) or by flow

cytometry (C).

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159

Figure 4.9. Expression of PaLoc promoters in a codY null mutant. The indicated strains were

grown in TY to stationary phase (24 h). Samples were fixed and analyzed using a plate reader

(A) or by flow cytometry (B). The bold arrow denotes a large number of cells saturated the

fluorescence detector during flow cytometry.

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CHAPTER V

Future Directions

Our studies have led to a deeper understanding of pathogenesis and cell division in C.

difficile. These insights have raised many new questions and directions for each project. Below,

we discuss these questions and strategies to answer them.

Chapter II

What are the functions of the Mld proteins? We showed the Mld proteins play a role

in cell division and morphology, but the exact functions of each Mld protein remain unknown.

Our general hypothesis is that they contribute to proper biogenesis of the peptidoglycan cell

wall. This hypothesis is based on the presence of a SPOR domain in MldA and the cell wall

defects seen when a mutant is grown in TY containing NaCl. In principle, the Mld proteins may

function as cell wall modifying enzymes directly (such as amidases, lytic transglycosylases, etc.).

However, none of the Mld proteins has a recognizable enzymatic domain and two of them are

predicted to contain coiled-coil motifs. These observations suggest the Mld proteins probably

affect peptidoglycan biogenesis indirectly by recruiting or activating enzymes that modify the

cell wall.

There are several ways in which these ideas could be pursued. First, the Mld proteins

(or their domains) could be purified and tested for peptidoglycan hydrolase activity. Second, a

pull-down may identify proteins associated with the Mld proteins. For example, anti-CFP

antibodies could be used to capture CFP-MldA and mass spectrometry methods could be used

to identify any proteins that co-immunoprecipitate with CFP-MldA. Analogous approaches have

been used to identify protein complexes involved in peptidoglycan assembly and turnover in

other bacteria (Kuhn et al., 2010; Sham et al., 2011). Putative interaction partners identified in

161

such a screen would need to be validated by asking whether those proteins also localize to the

midcell and by constructing mutants in the respective genes to determine whether there exhibit

morphological defects. Third, one could ask whether various known peptidoglycan synthases or

hydrolases mislocalize in an mldA::erm mutant. Finally, it would be worthwhile to determine

whether the dynamics of peptidoglycan synthesis or turnover is altered in an mldA::erm mutant.

This could be accomplished using newly introduced methods for labeling peptidoglycan by

incorporation of derivatives of D-alanine (Kuru et al., 2012; Kuru et al., 2015; Siegrist et al.,

2015; Siegrist et al., 2013). Pulse labeling with these compounds can reveal the rate and mode

of synthesis, while chase experiments provide information on degradation of the cell wall.

Genetic and biochemical analysis of the putative Mld complex. An important step

towards understanding how the Mld proteins work together during growth and division is to

determine where they reside in the cell, how they are localized to division sites, and whether

they form a complex.

For example, for a SPOR domain to function as such, it has to be in the extracellular

space, where it can bind peptidoglycan. Although prediction programs generally agree that

MldA has one transmembrane helix, they disagree on whether the N-terminus or the C-terminus

is extracellular. Because the SPOR domain is located near the C-terminus, we predict that MldA

should have an intracellular N-terminus and extracellular C-terminus. To confirm this topology

of MldA, we could use substituted-cysteine accessibility method (SCAM) (Bogdanov et al., 2005).

This method uses lipid-dependent regents to determine where a single cysteine in a protein is

located relative to the transmembrane. For this technique, the protein can only have one

cysteine (with the exception of a transmembrane cysteine). MldA has seven cysteine residues

so substitutions would be required. Alternatively, protease sensitivity has been used to

determine the topology of many transmembrane proteins, including, for example, germinant

162

receptors from Bacillus anthracis (Wilson et al., 2012). To apply this approach to MldA, C.

difficile cells producing CFP-MldA would be converted to spheroplasts and treated with

proteinase K in the presence or absence of Triton X-100 to disrupt membranes. Samples would

be analyzed by SDS-PAGE followed by Western blotting with anti-CFP antibodies. We predict

that CFP signal will only be lost if Triton X-100 is present during the protease step. Similar

approaches using CFP-MldB and CFP-MldC could be used to test the hypothesis that both of

these proteins are cytoplasmic.

To identify septal localization determinants in MldA, we would produce CFP fusions to

various deletion derivatives. A pilot experiment revealed a C-terminal truncation that removed

the predicted SPOR domain (CFP-MldA∆SPOR) still localizes to the midcell in an mldA null mutant

of C. difficile (data not shown). This finding is surprising because septal localization of SPOR

domain proteins is generally strongly dependent upon the presence of the SPOR domain

(Arends et al., 2010; Gerding et al., 2009; Möll and Thanbichler, 2009; Williams et al., 2013).

Perhaps MldA has multiple, independent septal targeting domains. Alternatively, the putative

SPOR domain might not really function as such.

The fact that localization of MldB and MldC depends upon MldA suggests these proteins

form a complex, as does the presence of predicted coiled-coil domains in MldA and MldB. The

immunoprecipitation approaches described should reveal whether all of the Mld proteins reside

in a complex. Alternative approaches include a bacterial two-hybrid system (Karimova et al.,

2005) for in vivo analyses or surface plasmon resonance for analyses using purified proteins or

protein domains (Chadsey et al., 1998).

163

Chapter III

Can mCherryOpt be used to monitor gene expression in the Syrian hamster model of

infection? There is much interest in studying gene expression in vivo, especially during the

course of a C. difficile infection. In Yersinia pseudotuberculosis virulence gene expression has

been visualized using promoter fusions to GFP and Ds2Red (Uliczka et al., 2011). For these

studies mice that had been infected with bacteria carrying appropriate reporter plasmids were

sacrificed, and cryosectioning was used to demonstrate specific gene expression in Peyer’s

Patches and mesenteric lymph nodes. In another example, a far red fluorescent protein called

Turbo-635 has been used to visualize Mycobacterium tuberculosis in the lungs of live mice

(Shcherbo et al., 2009). Based on these reports and a methods paper (Kong et al., 2005), we

think it is worth exploring whether mCherryOpt (or another RFP) can be used to visualize spatial

and temporal expression of C. difficile genes in the intestinal tract of hamsters or mice.

Questions that could be addressed include: Where do C. difficile spores reside in the

gastrointestinal tract? Where does germination occur following antibiotic treatment and what

are the kinetics for germination? When and where are the toxins expressed? Lastly, the use of

in vivo imaging of fluorescent proteins in the Syrian hamster may also help us address why C.

difficile mutants lacking the Mld proteins are attenuated.

Can mCherryOpt or another fluorescent protein be used in live C. difficile cells? One

of the major limitations of mCherryOpt is that chromophore maturation requires oxygen. But C.

difficile does not tolerate exposure to O2. Because of this, we have not been able to monitor

gene expression or aspects of C. difficile biology in live cells. The requirement for O2 precludes

use of mCherryOpt for studies of dynamic protein localization in live C. difficile cells. Moreover,

it is not possible to follow the fate of TcdA-ON cells: Do they stop producing toxin? Do they

lyse? Do they form spores?

164

Melville’s group was able to use a yellow fluorescent protein to study motility in

Clostridium perfringens by exposing the organism to air for about 20 min before returning it to

the anaerobic chamber (Hartman et al., 2011). Initial efforts to do this with C. difficile failed

because the organism lost about five logs of viability during a 5 min exposure to air (data not

shown), but it would be worthwhile to revisit this approach with different strains or in different

media/buffer conditions. The strain we used, JIR8094, is considered to be more O2-sensitive

than most other C. difficile. Another option would be newly discovered fluorescent proteins

that contain a bound flavin and therefore do not require O2 to become fluorescent (Mukherjee

and Schroeder, 2014). However, these are blue-green and not as bright at mCherryOpt.

Chapter IV

Determine the transcriptome of TcdA-ON and TcdA-OFF subpopulations of C. difficile.

It would be interesting to sort cells harboring a PtcdA::mCherryOpt reporter plasmid into TcdA-ON

and TcdA-OFF subpopulations, and analyze the transcriptomes by RNAseq. This approach

should reveal whether tcdB and tcdR are upregulated in the TcdA-ON subpopulation, as

expected. It also has the potential to identify any new genes that are co-regulated with tcdA;

these genes are likely to be pertinent to pathogenesis. Conversely, it seems likely that some

genes will be more highly expressed in the TcdA-OFF subpopulation. A prominent example

could be sporulation genes. Since sporulation is bistable in B. subtilis (Veening et al., 2005), we

anticipate that it may be a bistable phenotype in C. difficile as well. But whether cells that

produce toxin are distinct from those that sporulate is an open question. In addition, there are

other potential phenotypes which may be regulated distinctly from toxin production, including

motility and competence.

165

What is the relationship to toxin production to sporulation? As noted above, for

optimal transmission, C. difficile must form spores and produce toxins to cause diarrhea that

promotes release of the spores. These are two distinct cellular states, and we hypothesize that

toxin-producing cells do not sporulate. To test this, we could drive all C. difficile cells into a

toxin-producing state with our inducible TcdR construct and look for reduced sporulation. One

potential concern with interpretation of a TcdR overproduction experiment is that high levels of

TcdR might out-compete sporulation sigma factors for association with core RNA polymerase.

How are toxins exported? Little is known about how TcdA and TcdB are secreted from

the cell. Neither protein has a recognizable signal sequence, nor does the PaLoc encode an

obvious transport apparatus, with the possible exception of tcdE, which might (Carter et al.,

2011b; Dupuy et al., 2008; Matamouros et al., 2007) or might not (Cartman et al., 2012; Murray

et al., 2009) be a holin. Moreover, both toxins are about 300 kDa, and moving such enormous

proteins across the cytoplasmic membrane, through the peptidoglycan, and then across the S-

layer presents an interesting challenge.

One obvious experiment that must be done is to construct and characterize new null

mutants of tcdE. The issue of toxin export might also benefit from tagging the toxins with

fluorescent reports that enable them to be followed in real time. For example, one hypothesis is

that toxins are released by lysis. This idea can be tested by watching TcdA-ON cells to see if they

lyse. Tags that interfere with export might be lethal to C. difficile if they jam the export

apparatus. In this case, suppressor mutants might map to genes that encode factors involved in

toxin export and/or signal sequences that direct the toxins to the export machinery. These

approaches have proven successful for characterization of protein export in E. coli [reviewed in

(Beckwith, 2013)].

166

What controls bistability? Our data imply the alternative sigma TcdR is the master

regulator of bistability. Future efforts should be directed at determining whether tcdR

expression is bistable (as it is predicted to be), and understanding how tcdR expression is

regulated. Although TcdR clearly stimulates transcription of its own gene (Mani et al., 2002), the

relevant promoter(s) has yet to be definitively mapped. Two putative TcdR-dependent

promoters were identified at approximately -350 bp and -190 bp with respect to the ATG start

codon for tcdR (Mani et al., 2002). However, these promoters were inferred on the basis of

mobility shift assays using purified core RNA polymerase in combination with purified TcdR.

Curiously, neither putative promoter supported transcription in a purified system. Thus,

mapping transcriptional start sites at tcdR should be a high priority. If these promoters have

been correctly identified, they would give rise to mRNAs with unusually long leader sequences

that give rise to interesting possibilities for post-initiation regulation tcdR expression. Similarly,

the region upstream of tcdR has putative put poorly-characterized promoters recognized by SigA

and SigD [reviewed in (Bouillaut et al., 2015)].

Although binding sites for CodY and CcpA have been identified upstream of tcdR

(Antunes et al., 2011b; Dineen et al., 2007), the functional significance of these sites has yet to

be established. CodY is particularly interesting because expression of our PtcdA::mCherryOpt

reporter is greatly elevated in a subpopulation of cells from the codY::erm mutant. It is

surprising that we still observed a bistable phenotype despite the tremendous increase in

overall tcdA expression. This would suggest CodY only represses toxin expression in the TcdA-

ON subpopulation cells and has little or no effect in the TcdA-OFF cells. Is CodY production itself

bistable? In any event, our findings in the codY::erm mutant suggest a more complicated

regulation network than the current model.

167

Another open issue is the role of TcdC in toxin regulation. Conflicting reports claim it

has no effect on toxin production or serves as a negative regulator of toxin production (Bouvet

and Popoff, 2008; Carter et al., 2011b; Curry et al., 2007; Dupuy et al., 2008; Murray et al.,

2009). A useful starting point would be to construct a new null mutant. It would also be

interesting to evaluate the effect of ectopic expression of tcdC on expression of our

PtcdA::mCherryOpt reporter. Finally, it would be useful to know whether TcdC is produced in a

bistable manner and, if so, whether it is more abundant in the TcdA-ON or the TcdA-OFF

subpopulation of cells.

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References

Altschul, S.F., Wootton, J.C., Gertz, E.M., Agarwala, R., Morgulis, A., Schaffer, A.A., Yu, Y.K., 2005. Protein database searches using compositionally adjusted substitution matrices. Febs Journal 272, 5101-5109.

Andersen, J.B., Sternberg, C., Poulsen, L.K., Bjorn, S.P., Givskov, M., Molin, S., 1998. New unstable variants of green fluorescent protein for studies of transient gene expression in bacteria. Appl Environ Microbiol 64, 2240-2246.

Anderson, D.E., Gueiros-Filho, F.J., Erickson, H.P., 2004. Assembly dynamics of FtsZ rings in Bacillus subtilis and Escherichia coli and effects of FtsZ-regulating proteins. J Bacteriol 186, 5775-5781.

Anonymous, 2009. Genome of highly virulent Clostridium difficile analyzed. Expert Review of Anti-Infective Therapy 7, 1049-1049.

Antunes, A., Camiade, E., Monot, M., Courtois, E., Barbut, F., Sernova, N.V., Rodionov, D.A., Martin-Verstraete, I., Dupuy, B., 2012. Global transcriptional control by glucose and carbon regulator CcpA in Clostridium difficile. Nucleic Acids Res 40, 10701-10718.

Antunes, A., Martin-Verstraete, I., Dupuy, B., 2011a. CcpA-mediated repression of Clostridium difficile toxin gene expression. Mol Microbiol 79, 882-899.

Antunes, A., Martin-Verstraete, I., Dupuy, B., 2011b. CcpA-mediated repression of Clostridium difficile toxin gene expression. Mol Microbiol 79, 882-899.

Arends, S.J., Williams, K., Scott, R.J., Rolong, S., Popham, D.L., Weiss, D.S., 2010. Discovery and characterization of three new Escherichia coli septal ring proteins that contain a SPOR domain: DamX, DedD, and RlpA. J Bacteriol 192, 242-255.

Asai, K., Kawamura, F., Yoshikawa, H., Takahashi, H., 1995. Expression of Kina and Accumulation of Sigma(H) at the Onset of Sporulation in Bacillus subtilis. J Bacteriol 177, 6679-6683.

Aubry, A., Hussack, G., Chen, W.X., KuoLee, R., Twine, S.M., Fulton, K.M., Foote, S., Carrillo, C.D., Tanha, J., Logan, S.M., 2012. Modulation of Toxin Production by the Flagellar Regulon in Clostridium difficile. Infect Immun 80, 3521-3532.

Babcock, G.J., Broering, T.J., Hernandez, H.J., Mandell, R.B., Donahue, K., Boatright, N., Stack, A.M., Lowy, I., Graziano, R., Molrine, D., Ambrosino, D.M., Thomas, W.D., 2006. Human monoclonal antibodies directed against toxins A and B prevent Clostridium difficile-induced mortality in hamsters. Infect Immun 74, 6339-6347.

Bacci, S., Molbak, K., Kjeldsen, M.K., Olsen, K.E.P., 2011. Binary Toxin and Death after Clostridium difficile Infection. Emerging Infectious Diseases 17, 976-982.

Bacci, S., St-Martin, G., Olesen, B., Bruun, B., Olsen, K.E., Nielsen, E.M., Molbak, K., 2009. Outbreak of Clostridium difficile 027 in North Zealand, Denmark, 2008-2009. Euro Surveill 14.

Barbut, F., Decre, D., Lalande, V., Burghoffer, A., Noussair, L., Gigandon, A., Espinasse, F., Raskine, L., Robert, J., Mangeol, A., Branger, C., Petit, J.C., 2005. Clinical features of Clostridium difficile-associated diarrhoea due to binary toxin (actin-specific ADP-ribosyltransferase)-producing strains. J Med Microbiol 54, 181-185.

169

Barbut, F., Mastrantonio, P., Delmee, M., Brazier, J., Kuijper, E., Poxton, I., 2007. Prospective study of Clostridium difficile infections in Europe with phenotypic and genotypic characterisation of the isolates. Clinical Microbiology and Infection 13, 1048-1057.

Barbut, F., Petit, J.C., 2001. Epidemiology of Clostridium difficile-associated infections. Clinical Microbiology and Infection 7, 405-410.

Barra-Carrasco, J., Olguin-Araneda, V., Plaza-Garrido, A., Miranda-Cardenas, C., Cofre-Araneda, G., Pizarro-Guajardo, M., Sarker, M.R., Paredes-Sabja, D., 2013. The Clostridium difficile exosporium cysteine (CdeC)-rich protein is required for exosporium morphogenesis and coat assembly. J Bacteriol 195, 3863-3875.

Bartlett, J.G., 1994. Clostridium difficile - History of Its Role as an Enteric Pathogen and the Current State of Knowledge About the Organism. Clinical Infectious Diseases 18, S265-S272.

Bartlett, J.G., Chang, T.W., Gurwith, M., Gorbach, S.L., Onderdonk, A.B., 1978. Antibiotic-associated pseudomembranous colitis due to toxin-producing clostridia. N Engl J Med 298, 531-534.

Bartlett, J.G., Gerding, D.N., 2008. Clinical recognition and diagnosis of Clostridium difficile infection. Clin Infect Dis 46 Suppl 1, S12-18.

Bauer, M.P., Notermans, D.W., van Benthem, B.H.B., Brazier, J.S., Wilcox, M.H., Rupnik, M., Monnet, D.L., van Dissel, J.T., Kuijper, E.J., Grp, E.S., 2011. Clostridium difficile infection in Europe: a hospital-based survey. Lancet 377, 63-73.

Beckwith, J., 2013. The Sec-dependent pathway. Research in Microbiology 164, 497-504.

Belitsky, B.R., Sonenshein, A.L., 2013. Genome-wide identification of Bacillus subtilis CodY-binding sites at single-nucleotide resolution. Proc Natl Acad Sci U S A 110, 7026-7031.

Bera, A., Herbert, S., Jakob, A., Vollmer, W., Gotz, F., 2005. Why are pathogenic staphylococci so lysozyme resistant? The peptidoglycan O-acetyltransferase OatA is the major determinant for lysozyme resistance of Staphylococcus aureus. Mol Microbiol 55, 778-787.

Bogdanov, M., Zhang, W., Xie, J., Dowhan, W., 2005. Transmembrane protein topology mapping by the substituted cysteine accessibility method (SCAM (TM)): Application to lipid-specific membrane protein topogenesis. Methods 36, 148-171.

Borriello, S.P., 1990. The Influence of the Normal Flora on Clostridium difficile Colonization of the Gut. Annals of Medicine 22, 61-67.

Bouillaut, L., Dubois, T., Sonenshein, A.L., Dupuy, B., 2015. Integration of metabolism and virulence in Clostridium difficile. Res Microbiol 166, 375-383.

Bouillaut, L., McBride, S.M., Sorg, J.A., 2011. Genetic manipulation of Clostridium difficile. Curr Protoc Microbiol Chapter 9, Unit 9A 2.

Bouillaut, L., Self, W.T., Sonenshein, A.L., 2013. Proline-dependent regulation of Clostridium difficile Stickland metabolism. J Bacteriol 195, 844-854.

Bouvet, P.J.M., Popoff, M.R., 2008. Genetic Relatedness of Clostridium difficile Isolates from Various Origins Determined by Triple-Locus Sequence Analysis Based on Toxin Regulatory Genes tcdC, tcdR, and cdtR. Journal of Clinical Microbiology 46, 3703-3713.

170

Boyle, M.L., Ruth-Sahd, L.A., Zhou, Z., 2015. Fecal Microbiota Transplant to Treat Recurrent Clostridium difficile Infections. Critical Care Nurse 35, 51-63.

Braun, V., Hundsberger, T., Leukel, P., Sauerborn, M., von Eichel-Streiber, C., 1996. Definition of the single integration site of the pathogenicity locus in Clostridium difficile. Gene 181, 29-38.

Britton, R.A., Eichenberger, P., Gonzalez-Pastor, J.E., Fawcett, P., Monson, R., Losick, R., Grossman, A.D., 2002. Genome-wide analysis of the stationary-phase sigma factor (sigma-H) regulon of Bacillus subtilis. J Bacteriol 184, 4881-4890.

Brown, S., Santa Maria, J.P., Jr., Walker, S., 2013. Wall teichoic acids of gram-positive bacteria. Annu Rev Microbiol 67, 313-336.

Buffie, C.G., Jarchum, I., Equinda, M., Lipuma, L., Gobourne, A., Viale, A., Ubeda, C., Xavier, J., Pamer, E.G., 2012. Profound Alterations of Intestinal Microbiota following a Single Dose of Clindamycin Results in Sustained Susceptibility to Clostridium difficile-Induced Colitis. Infect Immun 80, 62-73.

Buss, J., Coltharp, C., Huang, T., Pohlmeyer, C., Wang, S.C., Hatem, C., Xiao, J., 2013. In vivo organization of the FtsZ-ring by ZapA and ZapB revealed by quantitative super-resolution microscopy. Mol Microbiol 89, 1099-1120.

Calabi, E., Ward, S., Wren, B., Paxton, T., Panico, M., Morris, H., Dell, A., Dougan, G., Fairweather, N., 2001. Molecular characterization of the surface layer proteins from Clostridium difficile. Mol Microbiol 40, 1187-1199.

Camorlingaponce, M., Torreslopez, J.F., Munozhernandez, O., 1985. Inhibition of Growth and Toxic Activity of Clostridium difficile by Normal Flora. Archivos De Investigacion Medica 16, 30-31.

Carlson, P.E., Walk, S.T., Bourgis, A.E.T., Liu, M.W., Kopliku, F., Lo, E., Young, V.B., Aronoff, D.M., Hanna, P.C., 2013. The relationship between phenotype, ribotype, and clinical disease in human Clostridium difficile isolates. Anaerobe 24, 109-116.

Carroll, K.C., Bartlett, J.G., 2011. Biology of Clostridium difficile: implications for epidemiology and diagnosis. Annu Rev Microbiol 65, 501-521.

Carter, G.P., Awad, M.M., Kelly, M.L., Rood, J.I., Lyras, D., 2011a. TcdB or not TcdB: a tale of two Clostridium difficile toxins. Future Microbiology 6, 121-123.

Carter, G.P., Douce, G.R., Govind, R., Howarth, P.M., Mackin, K.E., Spencer, J., Buckley, A.M., Antunes, A., Kotsanas, D., Jenkin, G.A., Dupuy, B., Rood, J.I., Lyras, D., 2011b. The anti-sigma factor TcdC modulates hypervirulence in an epidemic BI/NAP1/027 clinical isolate of Clostridium difficile. PLoS Pathog 7, e1002317.

Cartman, S.T., Kelly, M.L., Heeg, D., Heap, J.T., Minton, N.P., 2012. Precise manipulation of the Clostridium difficile chromosome reveals a lack of association between the tcdC genotype and toxin production. Appl Environ Microbiol 78, 4683-4690.

Cartman, S.T., Minton, N.P., 2010. A mariner-based transposon system for in vivo random mutagenesis of Clostridium difficile. Appl Environ Microbiol 76, 1103-1109.

171

Castilla-Llorente, V., Munoz-Espin, D., Villar, L., Salas, M., Meijer, W.J.J., 2006. Spo0A, the key transcriptional regulator for entrance into sporulation, is an inhibitor of DNA replication. Embo Journal 25, 3890-3899.

CDC, 2013. Antibiotic resistance threats in the United States, 2013. Centers for Disease Control and Prevention.

Chadsey, M.S., Karlinsey, J.E., Hughes, K.T., 1998. The flagellar anti-sigma factor FlgM actively dissociates Salmonella typhimurium sigma 28 RNA polymerase holoenzyme. Genes Dev 12, 3123-3136.

Chang, H.T., Krezolek, D., Johnson, S., Parada, J.P., Evans, C.T., Gerding, D.N., 2007. Onset of symptoms and time to diagnosis of Clostridium difficile - Associated disease following discharge from an acute care hospital. Infection Control and Hospital Epidemiology 28, 926-931.

Chang, T.W., Bartlett, J.G., Gorbach, S.L., Onderdonk, A.B., 1978. Clindamycin-Induced Enterocolitis in Hamsters as a Model of Pseudomembranous Colitis in Patients. Infect Immun 20, 526-529.

Chen, H.J., Tsai, T.K., Pan, S.C., Lin, J.S., Tseng, C.L., Shaw, G.C., 2010. The master transcription factor Spo0A is required for poly(3-hydroxybutyrate) (PHB) accumulation and expression of genes involved in PHB biosynthesis in Bacillus thuringiensis. FEMS Microbiol Lett 304, 74-81.

Chen, X.H., Katchar, K., Goldsmith, J.D., Nanthakumar, N., Cheknis, A., Gerding, D.N., Kelly, C.P., 2008. A Mouse Model of Clostridium difficile-Associated Disease. Gastroenterology 135, 1984-1992.

Chilcott, G.S., Hughes, K.T., 2000. Coupling of flagellar gene expression to flagellar assembly in Salmonella enterica serovar typhimurium and Escherichia coli. Microbiology and Molecular Biology Reviews 64, 694-+.

Chilton, C.H., Crowther, G.S., Freeman, J., Todhunter, S.L., Nicholson, S., Longshaw, C.M., Wilcox, M.H., 2013. Successful treatment of simulated Clostridium difficile infection in a human gut model by fidaxomicin first line and after vancomycin or metronidazole failure. J Antimicrob Chemother.

Choi, J.L., Kim, B.R., Kim, J.E., Woo, K.S., Kim, K.H., Kim, J.M., Lee, S.E., Han, J.Y., 2013. A case of Clostridium difficile bacteremia in a patient with loop ileostomy. Ann Lab Med 33, 200-202.

Chumbler, N.M., Farrow, M.A., Lapierre, L.A., Franklin, J.L., Haslam, D., Goldenring, J.R., Lacy, D.B., 2012. Clostridium difficile Toxin B Causes Epithelial Cell Necrosis through an Autoprocessing-Independent Mechanism. Plos Pathogens 8.

Cid, A., Juncal, A.R., Aguilera, A., Regueiro, B.J., Gonzalez, V., 1998. Clostridium difficile bacteremia in an immunocompetent child. J Clin Microbiol 36, 1167-1168.

Cloud, K.A., Dillard, J.P., 2004. Mutation of a single lytic transglycosylase causes aberrant septation and inhibits cell separation of Neisseria gonorrhoeae. J Bacteriol 186, 7811-7814.

Cohen, S.H., Gerding, D.N., Johnson, S., Kelly, C.P., Loo, V.G., McDonald, L.C., Pepin, J., Wilcox, M.H., 2010. Clinical Practice Guidelines for Clostridium difficile Infection in Adults: 2010 Update by the Society for Healthcare Epidemiology of America (SHEA) and the Infectious Diseases Society of America (IDSA). Infection Control and Hospital Epidemiology 31, 431-455.

172

Cohen, S.H., Tang, Y.J., Silva, J., 2000. Analysis of the pathogenicity locus in Clostridium difficile strains. Journal of Infectious Diseases 181, 659-663.

Colland, F., Rain, J.C., Gounon, P., Labigne, A., Legrain, P., De Reuse, H., 2001. Identification of the Helicobacter pylori anti-sigma(28) factor. Mol Microbiol 41, 477-487.

Collins, M.D., Lawson, P.A., Willems, A., Cordoba, J.J., Fernandez-Garayzabal, J., Garcia, P., Cai, J., Hippe, H., Farrow, J.A., 1994. The phylogeny of the genus Clostridium: proposal of five new genera and eleven new species combinations. Int J Syst Bacteriol 44, 812-826.

Comely, O.A., Miller, M.A., Louie, T.J., Crook, D.W., Gorbach, S.L., 2012. Treatment of First Recurrence of Clostridium difficile Infection: Fidaxomicin Versus Vancomycin. Clinical Infectious Diseases 55, S154-S161.

Cooperstock, M., Riegle, L., Woodruff, C.W., Onderdonk, A., 1983. Influence of Age, Sex, and Diet on Asymptomatic Colonization of Infants with Clostridium difficile. Journal of Clinical Microbiology 17, 830-833.

Cormack, B.P., Valdivia, R.H., Falkow, S., 1996. FACS-optimized mutants of the green fluorescent protein (GFP). Gene 173, 33-38.

Cosby, W.M., Zuber, P., 1997. Regulation of Bacillus subtilis sigma(H) (SpoOH) and AbrB in response to changes in external pH. J Bacteriol 179, 6778-6787.

Curry, S.R., Marsh, J.W., Muto, C.A., O'Leary, M.M., Pasculle, A.W., Harrison, L.H., 2007. tcdC genotypes associated with severe TcdC truncation in an epidemic clone and other strains of Clostridium difficile. J Clin Microbiol 45, 215-221.

Dai, K., Xu, Y., Lutkenhaus, J., 1993. Cloning and characterization of ftsN, an essential cell division gene in Escherichia coli isolated as a multicopy suppressor of ftsA12(Ts). J Bacteriol 175, 3790-3797.

Daniell, H.W., 2014. Increased Clostridium difficile Recurrences following Combined Proton Pump Inhibitor-Metronidazole Therapy. Infection Control and Hospital Epidemiology 35, 94-94.

Darling, A.E., Worden, P., Chapman, T.A., Chowdhury, P.R., Charles, I.G., Djordjevic, S.P., 2014. The genome of Clostridium difficile 5.3. Gut Pathogens 6.

Davies, H.A., Borriello, S.P., 1990. Detection of Capsule in Strains of Clostridium difficile of Varying Virulence and Toxigenicity. Microbial Pathogenesis 9, 141-146.

Dawson, L.F., Valiente, E., Faulds-Pain, A., Donahue, E.H., Wren, B.W., 2012. Characterisation of Clostridium difficile biofilm formation, a role for Spo0A. Plos One 7, e50527.

Deakin, L.J., Clare, S., Fagan, R.P., Dawson, L.F., Pickard, D.J., West, M.R., Wren, B.W., Fairweather, N.F., Dougan, G., Lawley, T.D., 2012. The Clostridium difficile spo0A gene is a persistence and transmission factor. Infect Immun 80, 2704-2711.

Delalla, F., Privitera, G., Ortisi, G., Rizzardini, G., Santoro, D., Pagano, A., Rinaldi, E., Scarpellini, P., 1989. 3rd-Generation Cephalosporins as a Risk Factor for Clostridium difficile-Associated Disease - a 4-Year Survey in a General-Hospital. Journal of Antimicrobial Chemotherapy 23, 623-631.

173

Delmee, M., Avesani, V., Delferriere, N., Burtonboy, G., 1990. Characterization of Flagella of Clostridium difficile and Their Role in Serogrouping Reactions. Journal of Clinical Microbiology 28, 2210-2214.

Dembek, M., Barquist, L., Boinett, C.J., Cain, A.K., Mayho, M., Lawley, T.D., Fairweather, N.F., Fagan, R.P., 2015. High-throughput analysis of gene essentiality and sporulation in Clostridium difficile. MBio 6, e02383.

Deutscher, J., Francke, C., Postma, P.W., 2008. How phosphotransferase system-related protein phosphorylation regulates carbohydrate metabolism in bacteria (vol 70, pg 939, 2006). Microbiology and Molecular Biology Reviews 72, 555-555.

Deutscher, J., Saier, M.H., 1983. Atp-Dependent Protein Kinase-Catalyzed Phosphorylation of a Seryl Residue in Hpr, a Phosphate Carrier Protein of the Phosphotransferase System in Streptococcus pyogenes. Proceedings of the National Academy of Sciences of the United States of America-Biological Sciences 80, 6790-6794.

Dineen, S.S., McBride, S.M., Sonenshein, A.L., 2010. Integration of Metabolism and Virulence by Clostridium difficile CodY. J Bacteriol 192, 5350-5362.

Dineen, S.S., Villapakkam, A.C., Nordman, J.T., Sonenshein, A.L., 2007. Repression of Clostridium difficile toxin gene expression by CodY. Mol Microbiol 66, 206-219.

Dingle, T., Wee, S., Mulvey, G.L., Greco, A., Kitova, E.N., Sun, J.X., Lin, S.J., Klassen, J.S., Palcic, M.M., Ng, K.K.S., Armstrong, G.D., 2008. Functional properties of the carboxy-terminal host cell-binding domains of the two toxins, TcdA and TcdB, expressed by Clostridium difficile. Glycobiology 18, 698-706.

Dixon, L.G., Seredick, S., Richer, M., Spiegelman, G.B., 2001. Developmental gene expression in Bacillus subtilis crsA47 mutants reveals glucose-activated control of the gene for the minor sigma factor sigma(H). J Bacteriol 183, 4814-4822.

Dominguez-Cuevas, P., Porcelli, I., Daniel, R.A., Errington, J., 2013. Differentiated roles for MreB-actin isologues and autolytic enzymes in Bacillus subtilis morphogenesis. Mol Microbiol 89, 1084-1098.

Dubberke, E.R., Reske, K.A., Yan, Y., Olsen, M.A., McDonald, L.C., Fraser, V.J., 2007. Clostridium difficile-associated disease in a setting of endemicity: Identification of novel risk factors. Clinical Infectious Diseases 45, 1543-1549.

Dubnau, D., Losick, R., 2006. Bistability in bacteria. Mol Microbiol 61, 564-572.

Dupuy, B., Govind, R., Antunes, A., Matamouros, S., 2008. Clostridium difficile toxin synthesis is negatively regulated by TcdC. J Med Microbiol 57, 685-689.

Dupuy, B., Sonenshein, A.L., 1998. Regulated transcription of Clostridium difficile toxin genes. Mol Microbiol 27, 107-120.

Eckert, C., Emirian, A., Le Monnier, A., Cathala, L., De Montclos, H., Goret, J., Berger, P., Petit, A., De Chevigny, A., Jean-Pierre, H., Nebbad, B., Camiade, S., Meckenstock, R., Lalande, V., Marchandin, H., Barbut, F., 2015. Prevalence and pathogenicity of binary toxin-positive Clostridium difficile strains that do not produce toxins A and B. New Microbes New Infect 3, 12-17.

174

Edwards, A.N., Suarez, J.M., McBride, S.M., 2013. Culturing and Maintaining Clostridium difficile in an Anaerobic Environment. Jove-Journal of Visualized Experiments.

El Meouche, I., Peltier, J., Monot, M., Soutourina, O., Pestel-Caron, M., Dupuy, B., Pons, J.L., 2013. Characterization of the SigD Regulon of C. difficile and Its Positive Control of Toxin Production through the Regulation of tcdR. Plos One 8.

Eyre, D.W., Fawley, W.N., Best, E.L., Griffiths, D., Stoesser, N.E., Crook, D.W., Peto, T.E.A., Walker, A.S., Wilcox, M.H., 2013. Comparison of Multilocus Variable-Number Tandem-Repeat Analysis and Whole-Genome Sequencing for Investigation of Clostridium difficile Transmission. Journal of Clinical Microbiology 51, 4141-4149.

Eyre, D.W., Walker, S., Wyllie, D., Dingle, K.E., Griffiths, D., Finney, J., O'Connor, L., Vaughan, A., Crook, D.W., Wilcox, M.H., Peto, T.E.A., Database, I.O.R., 2012. Predictors of First Recurrence of Clostridium difficile Infection: Implications for Initial Management. Clinical Infectious Diseases 55, S77-S87.

Fagan, R.P., Fairweather, N.F., 2011. Clostridium difficile has two parallel and essential Sec secretion systems. J Biol Chem 286, 27483-27493.

Fagan, R.P., Fairweather, N.F., 2014. Biogenesis and functions of bacterial S-layers. Nat Rev Microbiol 12, 211-222.

Farooq, P.D., Urrunaga, N.H., Tang, D.M., von Rosenvinge, E.C., 2015. Pseudomembranous colitis. Dis Mon 61, 181-206.

Fekety, R., McFarland, L.V., Surawicz, C.M., Greenberg, R.N., Elmer, G.W., Mulligan, M.E., 1997. Recurrent Clostridium difficile diarrhea: characteristics of and risk factors for patients enrolled in a prospective, randomized, double-blinded trial. Clin Infect Dis 24, 324-333.

Feldman, R.J., Kallich, M., Weinstein, M.P., 1995. Bacteremia due to Clostridium difficile: case report and review of extraintestinal C. difficile infections. Clin Infect Dis 20, 1560-1562.

Feltis, B.A., Wiesner, S.M., Kim, A.S., Erlandsen, S.L., Lyerly, D.L., Wilkins, T.D., Wells, C.L., 2000. Clostridium difficile toxins A and B can alter epithelial permeability and promote bacterial paracellular migration through HT-29 enterocytes. Shock 14, 629-634.

Finn, R.D., Bateman, A., Clements, J., Coggill, P., Eberhardt, R.Y., Eddy, S.R., Heger, A., Hetherington, K., Holm, L., Mistry, J., Sonnhammer, E.L., Tate, J., Punta, M., 2014. Pfam: the protein families database. Nucleic Acids Res 42, D222-230.

Forgetta, V., Oughton, M.T., Marquis, P., Brukner, I., Blanchette, R., Haub, K., Magrini, V., Mardis, E.R., Gerding, D.N., Loo, V.G., Miller, M.A., Mulvey, M.R., Rupnik, M., Dascal, A., Dewar, K., 2011. Fourteen-Genome Comparison Identifies DNA Markers for Severe-Disease-Associated Strains of Clostridium difficile. Journal of Clinical Microbiology 49, 2230-2238.

Formstone, A., Errington, J., 2005. A magnesium-dependent mreB null mutant: implications for the role of mreB in Bacillus subtilis. Mol Microbiol 55, 1646-1657.

Fujita, Y., 2009. Carbon Catabolite Control of the Metabolic Network in Bacillus subtilis. Bioscience Biotechnology and Biochemistry 73, 245-259.

Galli, E., Gerdes, K., 2010. Spatial resolution of two bacterial cell division proteins: ZapA recruits ZapB to the inner face of the Z-ring. Mol Microbiol 76, 1514-1526.

175

Genisyuerek, S., Papatheodorou, P., Guttenberg, G., Schubert, R., Benz, R., Aktories, K., 2011. Structural determinants for membrane insertion, pore formation and translocation of Clostridium difficile toxin B. Mol Microbiol 79, 1643-1654.

George, R.H., Symonds, J.M., Dimock, F., Brown, J.D., Arabi, Y., Shinagawa, N., Keighley, M.R., Alexander-Williams, J., Burdon, D.W., 1978. Identification of Clostridium difficile as a cause of pseudomembranous colitis. British Medical Journal 1, 695-695.

Gerding, D.N., 2004. Clindamycin, cephalosporins, fluoroquinolones, and Clostridium difficile-associated diarrhea: This is an antimicrobial resistance problem. Clinical Infectious Diseases 38, 646-648.

Gerding, M.A., Liu, B., Bendezu, F.O., Hale, C.A., Bernhardt, T.G., de Boer, P.A., 2009. Self-enhanced accumulation of FtsN at Division Sites and Roles for Other Proteins with a SPOR domain (DamX, DedD, and RlpA) in Escherichia coli cell constriction. J Bacteriol 191, 7383-7401.

Gerhard, R., Frenzel, E., Goy, S., Olling, A., 2013. Cellular uptake of Clostridium difficile TcdA and truncated TcdA lacking the receptor binding domain. J Med Microbiol 62, 1414-1422.

Gerhard, R., Nottrott, S., Schoentaube, J., Tatge, H., Olling, A., Just, I., 2008. Glucosylation of Rho GTPases by Clostridium difficile toxin A triggers apoptosis in intestinal epithelial cells. J Med Microbiol 57, 765-770.

Geric, B., Carman, R.J., Rupnik, M., Genheimer, C.W., Sambol, S.P., Lyerly, D.M., Gerding, D.N., Johnson, S., 2006. Binary toxin-producing, large clostridial toxin-negative Clostridium difficile strains are enterotoxic but do not cause disease in hamsters. J Infect Dis 193, 1143-1150.

Geric, B., Johnson, S., Gerding, D.N., Grabnar, M., Rupnik, M., 2003. Frequency of binary toxin genes among Clostridium difficile strains that do not produce large clostridial toxins. J Clin Microbiol 41, 5227-5232.

Gode-Potratz, C.J., Kustusch, R.J., Breheny, P.J., Weiss, D.S., McCarter, L.L., 2011. Surface sensing in Vibrio parahaemolyticus triggers a programme of gene expression that promotes colonization and virulence. Mol Microbiol 79, 240-263.

Goldstein, B.A., Fabiano, L., Washington, P.S., 2005. Phonological skills in predominantly English-speaking, predominantly Spanish-speaking, and Spanish-English bilingual children. Lang Speech Hear Serv Sch 36, 201-218.

Govind, R., Dupuy, B., 2012. Secretion of Clostridium difficile toxins A and B requires the holin-like protein TcdE. PLoS Pathog 8, e1002727.

Greathouse, K.L., Harris, C.C., Bultman, S.J., 2015. Dysfunctional Families: Clostridium scindens and Secondary Bile Acids Inhibit the Growth of Clostridium difficile. Cell Metabolism 21, 9-10.

Greco, A., Ho, J.G.S., Lin, S.J., Palcic, M.M., Rupnik, M., Ng, K.K.S., 2006. Carbohydrate recognition by Clostridium difficile toxin A. Nature Structural & Molecular Biology 13, 460-461.

Guedon, E., Serror, P., Ehrlich, S.D., Renault, P., Delorme, C., 2001. Pleiotropic transcriptional repressor CodY senses the intracellular pool of branched-chain amino acids in Lactococcus lactis. Mol Microbiol 40, 1227-1239.

Gueiros-Filho, F.J., Losick, R., 2002. A widely conserved bacterial cell division protein that promotes assembly of the tubulin-like protein FtsZ. Genes Dev 16, 2544-2556.

176

Hadjifrangiskou, M., Chen, Y.H., Koehler, T.A., 2007. The alternative sigma factor sigma(H) is required for toxin gene expression by Bacillus anthracis. J Bacteriol 189, 1874-1883.

Hafiz, S., Oakley, C.L., 1976. Clostridium difficile - Isolation and Characteristics. J Med Microbiol 9, 129-&.

Hahn, J., Roggiani, M., Dubnau, D., 1995. The major role of Spo0A in genetic competence is to downregulate abrB, an essential competence gene. J Bacteriol 177, 3601-3605.

Hall, I.C., O'Toole, E., 1935. Intestinal flora in new-born infants with a description of a new pathogenic anaerobe, Bacillus difficilis. American Journal of Diseases of Children 49, 390-402.

Hamon, M.A., Lazazzera, B.A., 2001. The sporulation transcription factor Spo0A is required for biofilm development in Bacillus subtilis. Mol Microbiol 42, 1199-1209.

Hamon, M.A., Stanley, N.R., Britton, R.A., Grossman, A.D., Lazazzera, B.A., 2004. Identification of AbrB-regulated genes involved in biofilm formation by Bacillus subtilis. Mol Microbiol 52, 847-860.

Han, S.J., Cheon, J.W., Park, J.B., Kim, K.Y., Kim, Y.K., Kim, B.S., Han, J., Lee, C.H., Kim, H.G., Jung, J.T., Kwon, J.G., Kim, E.Y., 2009. Clinical factors for severity and relapse of pseudomembranous colitis. Journal of Gastroenterology and Hepatology 24, A71-A71.

Hartman, A.H., Liu, H.L., Melville, S.B., 2011. Construction and Characterization of a Lactose-Inducible Promoter System for Controlled Gene Expression in Clostridium perfringens. Appl Environ Microbiol 77, 471-478.

Hartmann, B., Just, I., Genth, H., 2003. Intracellular stability of rho proteins glucosylated by Clostridium difficile Toxin B. Naunyn-Schmiedebergs Archives of Pharmacology 367, R55-R55.

Hatheway, C.L., 1990. Toxigenic clostridia. Clin Microbiol Rev 3, 66-98.

He, M., Sebaihia, M., Lawley, T.D., Stabler, R.A., Dawson, L.F., Martin, M.J., Holt, K.E., Seth-Smith, H.M.B., Quail, M.A., Rance, R., Brooks, K., Churcher, C., Harris, D., Bentley, S.D., Burrows, C., Clark, L., Corton, C., Murray, V., Rose, G., Thurston, S., van Tonder, A., Walker, D., Wren, B.W., Dougan, G., Parkhill, J., 2010. Evolutionary dynamics of Clostridium difficile over short and long time scales. Proc Natl Acad Sci U S A 107, 7527-7532.

Heap, J.T., Kuehne, S.A., Ehsaan, M., Cartman, S.T., Cooksley, C.M., Scott, J.C., Minton, N.P., 2010. The ClosTron: Mutagenesis in Clostridium refined and streamlined. J Microbiol Methods 80, 49-55.

Heap, J.T., Pennington, O.J., Cartman, S.T., Minton, N.P., 2009. A modular system for Clostridium shuttle plasmids. J Microbiol Methods 78, 79-85.

Heidrich, C., Templin, M.F., Ursinus, A., Merdanovic, M., Berger, J., Schwarz, H., de Pedro, M.A., Höltje, J.V., 2001. Involvement of N-acetylmuramyl-L-alanine amidases in cell separation and antibiotic-induced autolysis of Escherichia coli. Mol Microbiol 41, 167-178.

Heim, R., Prasher, D.C., Tsien, R.Y., 1994. Wavelength mutations and posttranslational autoxidation of green fluorescent protein. Proc Natl Acad Sci U S A 91, 12501-12504.

Heine, K., Barth, H., 2006. Role of Rho-GTPases in Clostridium difficile toxin induced cell death in macrophage-like cells. Naunyn-Schmiedebergs Archives of Pharmacology 372, 120-120.

177

Helmann, J.D., Chamberlin, M.J., 1987. DNA-Sequence Analysis Suggests That Expression of Flagellar and Chemotaxis Genes in Escherichia coli and Salmonella typhimurium Is Controlled by an Alternative Sigma-Factor. Proc Natl Acad Sci U S A 84, 6422-6424.

Hemminger, J., Balada-Llasat, J.M., Raczkowski, M., Buckosh, M., Pancholi, P., 2011. Two case reports of Clostridium difficile bacteremia, one with the epidemic NAP-1 strain. Infection 39, 371-373.

Ho, T.D., Ellermeier, C.D., 2011. PrsW is required for colonization, resistance to antimicrobial peptides, and expression of extracytoplasmic function sigma factors in Clostridium difficile. Infect Immun 79, 3229-3238.

Ho, T.D., Williams, K.B., Chen, Y., Helm, R.F., Popham, D.L., Ellermeier, C.D., 2014. Clostridium difficile extracytoplasmic function sigma factor sigmaV regulates lysozyme resistance and is necessary for pathogenesis in the hamster model of infection. Infect Immun 82, 2345-2355.

Horstmann, N., Seidel, G., Aung-Hilbrich, L.M., Hillen, W., 2007. Residues His-15 and Arg-17 of HPr participate differently in catabolite signal processing via CcpA. Journal of Biological Chemistry 282, 1175-1182.

Hsiao, C.H., Ueno, N., Shao, J.Q., Schroeder, K.R., Moore, K.C., Donelson, J.E., Wilson, M.E., 2011. The effects of macrophage source on the mechanism of phagocytosis and intracellular survival of Leishmania. Microbes Infect 13, 1033-1044.

Hundsberger, T., Braun, V., Weidmann, M., Leukel, P., Sauerborn, M., von Eichel-Streiber, C., 1997. Transcription analysis of the genes tcdA-E of the pathogenicity locus of Clostridium difficile. Eur J Biochem 244, 735-742.

Indra, A., Lassnig, H., Baliko, N., Much, P., Fiedler, A., Huhulescu, S., Allerberger, F., 2009. Clostridium difficile: a new zoonotic agent? Wien Klin Wochenschr 121, 91-95.

Jackson, S., Calos, M., Myers, A., Self, W.T., 2006. Analysis of proline reduction in the nosocomial pathogen Clostridium difficile. J Bacteriol 188, 8487-8495.

Jagai, J., Naumova, E., 2009. Clostridium difficile-associated Disease in the Elderly, United States. Emerging Infectious Diseases 15, 343-344.

Jank, T., Giesemann, T., Aktories, K., 2007a. Clostridium difficile glucosyltransferase toxin B-essential amino acids for substrate binding. J Biol Chem 282, 35222-35231.

Jank, T., Giesemann, T., Aktories, K., 2007b. Rho-glucosylating Clostridium difficile toxins A and B: new insights into structure and function. Glycobiology 17, 15R-22R.

Johnson, B.P., Jensen, B.J., Ransom, E.M., Heinemann, K.A., Vannatta, K.M., Egland, K.A., Egland, P.G., 2009. Interspecies signaling between Veillonella atypica and Streptococcus gordonii requires the transcription factor CcpA. J Bacteriol 191, 5563-5565.

Johnson, E.A., 1999. Clostridial toxins as therapeutic agents: benefits of nature's most toxic proteins. Annu Rev Microbiol 53, 551-575.

Johnson, J.E., Lackner, L.L., Hale, C.A., de Boer, P.A., 2004. ZipA is required for targeting of DMinC/DicB, but not DMinC/MinD, complexes to septal ring assemblies in Escherichia coli. J Bacteriol 186, 2418-2429.

178

Jones, A.M., Kuijper, E.J., Wilcox, M.H., 2013. Clostridium difficile: a European perspective. J Infect 66, 115-128.

Just, I., Fritz, G., Aktories, K., Giry, M., Popoff, M.R., Boquet, P., Hegenbarth, S., Voneichelstreiber, C., 1994. Clostridium difficile Toxin-B Acts on the Gtp-Binding Protein-Rho. Journal of Biological Chemistry 269, 10706-10712.

Just, I., Selzer, J., Voneichelstreiber, C., Aktories, K., 1995a. The Low-Molecular-Mass Gtp-Binding Protein-Rho Is Affected by Toxin-a from Clostridium difficile. Journal of Clinical Investigation 95, 1026-1031.

Just, I., Selzer, J., Wilm, M., Voneichelstreiber, C., Mann, M., Aktories, K., 1995b. Glucosylation of Rho-Proteins by Clostridium difficile Toxin-B. Nature 375, 500-503.

Just, I., Wilm, M., Selzer, J., Rex, G., Voneichelstreiber, C., Mann, M., Aktories, K., 1995c. The Enterotoxin from Clostridium difficile (Toxa) Monoglucosylates the Rho-Proteins. Journal of Biological Chemistry 270, 13932-13936.

Kaplan, E.L., Meier, P., 1958. Nonparametric estimation from incomplete observations. J Am Stat Assoc 53, 457-481.

Karimova, G., Dautin, N., Ladant, D., 2005. Interaction network among Escherichia coli membrane proteins involved in cell division as revealed by bacterial two-hybrid analysis. J Bacteriol 187, 2233-2243.

Karlsson, S., Burman, L.G., Akerlund, T., 1999. Suppression of toxin production in Clostridium difficile VPI 10463 by amino acids. Microbiology-Uk 145, 1683-1693.

Karlsson, S., Burman, L.G., Akerlund, T., 2008. Induction of toxins in Clostridium difficile is associated with dramatic changes of its metabolism. Microbiology-Sgm 154, 3430-3436.

Karlsson, S., Dupuy, B., Mukherjee, K., Norin, E., Burman, L.G., Akerlund, T., 2003. Expression of Clostridium difficile toxins A and B and their sigma factor TcdD is controlled by temperature. Infect Immun 71, 1784-1793.

Karlsson, S., Lindberg, A., Norin, E., Burman, L.G., Akerlund, T., 2000. Toxins, butyric acid, and other short-chain fatty acids are coordinately expressed and down-regulated by cysteine in Clostridium difficile. Infect Immun 68, 5881-5888.

Kato, H., Kita, H., Karasawa, T., Maegawa, T., Koino, Y., Takakuwa, H., Saikai, T., Kobayashi, K., Yamagishi, T., Nakamura, S., 2001. Colonisation and transmission of Clostridium difficile in healthy individuals examined by PCR ribotyping and pulsed-field gel electrophoresis. J Med Microbiol 50, 720-727.

Kaufman, E., Liska, D., Rubinshteyn, V., Nandakumar, G., 2013. Clostridium difficile bacteremia. Surg Infect (Larchmt) 14, 559-560.

Keel, K., Brazier, J.S., Post, K.W., Weese, S., Songer, J.G., 2007. Prevalence of PCR ribotypes among Clostridium difficile isolates from pigs, calves, and other species. Journal of Clinical Microbiology 45, 1963-1964.

Kelly, C.P., Becker, S., Linevsky, J.K., Joshi, M.A., Okeane, J.C., Dickey, B.F., Lamont, J.T., Pothoulakis, C., 1994. Neutrophil Recruitment in Clostridium difficile Toxin-a Enteritis in the Rabbit. Journal of Clinical Investigation 93, 1257-1265.

179

Kelly, C.P., LaMont, J.T., 2008. Clostridium difficile--more difficult than ever. N Engl J Med 359, 1932-1940.

Kelly, C.R., Ihunnah, C., Fischer, M., Khoruts, A., Surawicz, C., Afzali, A., Aroniadis, O., Barto, A., Borody, T., Giovanelli, A., Gordon, S., Gluck, M., Hohmann, E.L., Kao, D., Kao, J.Y., McQuillen, D.P., Mellow, M., Rank, K.M., Rao, K., Ray, A., Schwartz, M.A., Singh, N., Stollman, N., Suskind, D.L., Vindigni, S.M., Youngster, I., Brandt, L., 2014. Fecal Microbiota Transplant for Treatment of Clostridium difficile Infection in Immunocompromised Patients. American Journal of Gastroenterology 109, 1065-1071.

Ketley, J.M., Haslam, S.C., Mitchell, T.J., Stephen, J., Candy, D.C.A., Burdon, D.W., 1984. Production and Release of Toxins a and B by Clostridium difficile. J Med Microbiol 18, 385-391.

Khan, M.Y., Hall, W.H., 1966. Staphylococcal enterocolitis--treatment with oral vancomycin. Annals of Internal Medicine 65, 1-8.

Kim, K.H., Fekety, R., Batts, D.H., Brown, D., Cudmore, M., Silva, J., Jr., Waters, D., 1981. Isolation of Clostridium difficile from the environment and contacts of patients with antibiotic-associated colitis. J Infect Dis 143, 42-50.

Kong, Y., Akin, A.R., Francis, K.P., Zhang, N., Troy, T.L., Xie, H., Rao, J., Cirillo, S.L.G., Cirillo, J.D., 2005. Whole-Body Imaging of Infection Using Fluorescence, Curr Protoc Microbiol. John Wiley & Sons, Inc.

Kuehne, S.A., Cartman, S.T., Heap, J.T., Kelly, M.L., Cockayne, A., Minton, N.P., 2010. The role of toxin A and toxin B in Clostridium difficile infection. Nature 467, 711-U797.

Kuehne, S.A., Collery, M.M., Kelly, M.L., Cartman, S.T., Cockayne, A., Minton, N.P., 2014. Importance of Toxin A, Toxin B, and CDT in Virulence of an Epidemic Clostridium difficile Strain. Journal of Infectious Diseases 209, 83-86.

Kuhn, J., Briegel, A., Morschel, E., Kahnt, J., Leser, K., Wick, S., Jensen, G.J., Thanbichler, M., 2010. Bactofilins, a ubiquitous class of cytoskeletal proteins mediating polar localization of a cell wall synthase in Caulobacter crescentus. Embo Journal 29, 327-339.

Kuipers, E.J., Surawicz, C.M., 2008. Clostridium difficile infection. Lancet 371, 1486-1488.

Kuru, E., Hughes, H.V., Brown, P.J., Hall, E., Tekkam, S., Cava, F., de Pedro, M.A., Brun, Y.V., VanNieuwenhze, M.S., 2012. In Situ Probing of Newly Synthesized Peptidoglycan in Live Bacteria with Fluorescent D-Amino Acids. Angewandte Chemie-International Edition 51, 12519-12523.

Kuru, E., Tekkam, S., Hall, E., Brun, Y.V., Van Nieuwenhze, M.S., 2015. Synthesis of fluorescent D-amino acids and their use for probing peptidoglycan synthesis and bacterial growth in situ. Nature Protocols 10, 33-52.

Kyne, L., Merry, C., O'Connell, B., Kelly, A., Keane, C., O'Neill, D., 1999. Factors associated with prolonged symptoms and severe disease due to Clostridium difficile. Age and Ageing 28, 107-113.

Larson, H.E., Price, A.B., Honour, P., Borriello, S.P., 1978. Clostridium difficile and the aetiology of pseudomembranous colitis. Lancet 1, 1063-1066.

180

Lawley, T.D., Clare, S., Deakin, L.J., Goulding, D., Yen, J.L., Raisen, C., Brandt, C., Lovell, J., Cooke, F., Clark, T.G., Dougan, G., 2010. Use of Purified Clostridium difficile Spores To Facilitate Evaluation of Health Care Disinfection Regimens. Appl Environ Microbiol 76, 6895-6900.

Lawley, T.D., Clare, S., Walker, A.W., Goulding, D., Stabler, R.A., Croucher, N., Mastroeni, P., Scott, P., Raisen, C., Mottram, L., Fairweather, N.F., Wren, B.W., Parkhill, J., Dougan, G., 2009. Antibiotic treatment of Clostridium difficile carrier mice triggers a supershedder state, spore-mediated transmission, and severe disease in immunocompromised hosts. Infect Immun 77, 3661-3669.

Lequette, Y., Greenberg, E.P., 2005. Timing and localization of rhamnolipid synthesis gene expression in Pseudomonas aeruginosa biofilms. J Bacteriol 187, 37-44.

Levett, P.N., 1984. Detection of Clostridium difficile in faeces by direct gas liquid chromatography. J Clin Pathol 37, 117-119.

Libby, D.B., Bearman, G., 2009. Bacteremia due to Clostridium difficile--review of the literature. Int J Infect Dis 13, e305-309.

Louie, T.J., Miller, M.A., Mullane, K.M., Weiss, K., Lentnek, A., Golan, Y., Gorbach, S., Sears, P., Shue, Y.K., Grp, O.-.-C.S., 2011. Fidaxomicin versus Vancomycin for Clostridium difficile Infection. New England Journal of Medicine 364, 422-431.

Low, H.H., Moncrieffe, M.C., Lowe, J., 2004. The crystal structure of ZapA and its modulation of FtsZ polymerisation. J Mol Biol 341, 839-852.

Lowy, I., Molrine, D.C., Leav, B.A., Blair, B.M., Baxter, R., Gerding, D.N., Nichol, G., Thomas, W.D., Leney, M., Sloan, S., Hay, C.A., Ambrosino, D.M., 2010. Treatment with Monoclonal Antibodies against Clostridium difficile Toxins. New England Journal of Medicine 362, 197-205.

Lupas, A., 1996. Prediction and analysis of coiled-coil structures. Methods Enzymol 266, 513-525.

Lyerly, D.M., Lockwood, D.E., Richardson, S.H., Wilkins, T.D., 1982. Biological-Activities of Toxin-A and Toxin-B of Clostridium difficile. Infect Immun 35, 1147-1150.

Lyerly, D.M., Saum, K.E., Macdonald, D.K., Wilkins, T.D., 1985. Effects of Clostridium difficile Toxins Given Intragastrically to Animals. Infect Immun 47, 349-352.

Lyngstadaas, A., Lobner-Olesen, A., Boye, E., 1995. Characterization of three genes in the dam-containing operon of Escherichia coli. Mol Gen Genet 247, 546-554.

Lyras, D., O'Connor, J.R., Howarth, P.M., Sambol, S.P., Carter, G.P., Phumoonna, T., Poon, R., Adams, V., Vedantam, G., Johnson, S., Gerding, D.N., Rood, J.I., 2009. Toxin B is essential for virulence of Clostridium difficile. Nature 458, 1176-1179.

Maamar, H., Raj, A., Dubnau, D., 2007. Noise in gene expression determines cell fate in Bacillus subtilis. Science 317, 526-529.

Mackin, K.E., Carter, G.P., Howarth, P., Rood, J.I., Lyras, D., 2013. Spo0A Differentially Regulates Toxin Production in Evolutionarily Diverse Strains of Clostridium difficile. Plos One 8.

181

Mahr, K., Esteban, C.D., Hillen, W., Titgemeyer, F., Perez-Martinez, G., 2002. Cross communication between components of carbon catabolite repression of Lactobacillus casei and Bacillus megaterium. Journal of Molecular Microbiology and Biotechnology 4, 489-494.

Majerczyk, C.D., Sadykov, M.R., Luong, T.T., Lee, C., Somerville, G.A., Sonenshein, A.L., 2008. Staphylococcus aureus CodY negatively regulates virulence gene expression. J Bacteriol 190, 2257-2265.

Mani, N., Dupuy, B., 2001. Regulation of toxin synthesis in Clostridium difficile by an alternative RNA polymerase sigma factor. Proc Natl Acad Sci U S A 98, 5844-5849.

Mani, N., Lyras, D., Barroso, L., Howarth, P., Wilkins, T., Rood, J.I., Sonenshein, A.L., Dupuy, B., 2002. Environmental response and autoregulation of Clostridium difficile TxeR, a sigma factor for toxin gene expression. J Bacteriol 184, 5971-5978.

Margolin, W., 2012. The price of tags in protein localization studies. J Bacteriol 194, 6369-6371.

Marquez, L.M., Helmann, J.D., Ferrari, E., Parker, H.M., Ordal, G.W., Chamberlin, M.J., 1990. Studies of Sigma-D-Dependent Functions in Bacillus subtilis. J Bacteriol 172, 3435-3443.

Martin, M.J., Clare, S., Goulding, D., Faulds-Pain, A., Barquist, L., Browne, H.P., Pettit, L., Dougan, G., Lawley, T.D., Wren, B.W., 2013. The agr Locus Regulates Virulence and Colonization Genes in Clostridium difficile 027. J Bacteriol 195, 3672-3681.

Matamouros, S., England, P., Dupuy, B., 2007. Clostridium difficile toxin expression is inhibited by the novel regulator TcdC. Mol Microbiol 64, 1274-1288.

McBee, R.H., 1960. INTESTINAL FLORA OF SOME ANTARCTIC BIRDS AND MAMMALS. J Bacteriol 79, 311-312.

McBride, S.M., Sonenshein, A.L., 2011. Identification of a genetic locus responsible for antimicrobial peptide resistance in Clostridium difficile. Infect Immun 79, 167-176.

McFarland, L.V., Elmer, G.W., Surawicz, C.M., 2002. Breaking the cycle: Treatment strategies for 163 cases of recurrent Clostridium difficile disease. American Journal of Gastroenterology 97, 1769-1775.

McKee, R.W., Mangalea, M.R., Purcell, E.B., Borchardt, E.K., Tamayo, R., 2013. The Second Messenger Cyclic Di-GMP Regulates Clostridium difficile Toxin Production by Controlling Expression of sigD. J Bacteriol 195, 5174-5185.

Meisner, J., Montero Llopis, P., Sham, L.T., Garner, E., Bernhardt, T.G., Rudner, D.Z., 2013. FtsEX is required for CwlO peptidoglycan hydrolase activity during cell wall elongation in Bacillus subtilis. Mol Microbiol 89, 1069-1083.

Mercer, K.L., Weiss, D.S., 2002. The Escherichia coli cell division protein FtsW is required to recruit its cognate transpeptidase, FtsI (PBP3), to the division site. J Bacteriol 184, 904-912.

Merrigan, M., Venugopal, A., Mallozzi, M., Roxas, B., Viswanathan, V.K., Johnson, S., Gerding, D.N., Vedantam, G., 2010. Human Hypervirulent Clostridium difficile Strains Exhibit Increased Sporulation as Well as Robust Toxin Production. J Bacteriol 192, 4904-4911.

Migocki, M.D., Freeman, M.K., Wake, R.G., Harry, E.J., 2002. The Min system is not required for precise placement of the midcell Z ring in Bacillus subtilis. EMBO Rep 3, 1163-1167.

182

Mirel, D.B., Chamberlin, M.J., 1989. The Bacillus subtilis Flagellin Gene (Hag) Is Transcribed by the Sigma-28 Form of Rna-Polymerase. J Bacteriol 171, 3095-3101.

Mishima, M., Shida, T., Yabuki, K., Kato, K., Sekiguchi, J., Kojima, C., 2005. Solution structure of the peptidoglycan binding domain of Bacillus subtilis cell wall lytic enzyme CwlC: characterization of the sporulation-related repeats by NMR. Biochemistry 44, 10153-10163.

Mitas, L., Svaton, R., Skricka, T., Kala, Z., Penka, I., Hanslianov, M., Grolich, T., Polak, P., Hlavsa, J., 2012. Surgical treatment of Clostridium colitides. Acta Chir Iugosl 59, 63-69.

Möll, A., Thanbichler, M., 2009. FtsN-like proteins are conserved components of the cell division machinery in proteobacteria. Mol Microbiol 72, 1037-1053.

Moncrief, J.S., Barroso, L.A., Wilkins, T.D., 1997. Positive regulation of Clostridium difficile toxins. Infect Immun 65, 1105-1108.

Monot, M., Boursaux-Eude, C., Thibonnier, M., Vallenet, D., Moszer, I., Medigue, C., Martin-Verstraete, I., Dupuy, B., 2011. Reannotation of the genome sequence of Clostridium difficile strain 630. J Med Microbiol 60, 1193-1199.

Moore, P., Kyne, L., Martin, A., Solomon, K., 2013. Germination efficiency of clinical Clostridium difficile spores and correlation with ribotype, disease severity and therapy failure. J Med Microbiol 62, 1405-1413.

Mukherjee, A., Schroeder, C.M., 2014. Flavin-based fluorescent proteins: emerging paradigms in biological imaging. Curr Opin Biotechnol 31C, 16-23.

Murray, R., Boyd, D., Levett, P.N., Mulvey, M.R., Alfa, M.J., 2009. Truncation in the tcdC region of the Clostridium difficile PathLoc of clinical isolates does not predict increased biological activity of Toxin B or Toxin A. Bmc Infectious Diseases 9.

Murray, T., Popham, D.L., Setlow, P., 1998. Bacillus subtilis cells lacking penicillin-binding protein 1 require increased levels of divalent cations for growth. J Bacteriol 180, 4555-4563.

Na, X., Kim, H., Moyer, M.P., Pothoulakis, C., LaMont, J.T., 2008. gp96 is a human colonocyte plasma membrane binding protein for Clostridium difficile toxin A. Infect Immun 76, 2862-2871.

Nakamura, S., Mikawa, M., Nakashio, S., Takabatake, M., Okado, I., Yamakawa, K., Serikawa, T., Okumura, S., Nishida, S., 1981. Isolation of Clostridium difficile from the Feces and the Antibody in Sera of Young and Elderly Adults. Microbiology and Immunology 25, 345-351.

Nakamura, Y., Gojobori, T., Ikemura, T., 2000. Codon usage tabulated from international DNA sequence databases: status for the year 2000. Nucleic Acids Res 28, 292.

Nelson, R.L., Kelsey, P., Leeman, H., Meardon, N., Patel, H., Paul, K., Rees, R., Taylor, B., Wood, E., Malakun, R., 2011. Antibiotic treatment for Clostridium difficile-associated diarrhea in adults. Cochrane Database Syst Rev, CD004610.

Nusrat, A., Turner, J.R., Parkos, C.A., vonEichelStreiber, C., Anderson, J.M., Carnes, D.K., Madara, J.L., 1996. Rho uncoupling by Clostridium difficile toxins A and B displaces tight junction associated proteins from the membrane of polarized epithelial cells. Molecular Biology of the Cell 7, 3533-3533.

183

O'Connor, J.R., Lyras, D., Farrow, K.A., Adams, V., Powell, D.R., Hinds, J., Cheung, J.K., Rood, J.I., 2006. Construction and analysis of chromosomal Clostridium difficile mutants. Mol Microbiol 61, 1335-1351.

O'Horo, J.C., Jindai, K., Kunzer, B., Safdar, N., 2013. Treatment of recurrent Clostridium difficile infection: a systematic review. Infection.

Olling, A., Goy, S., Tatge, H., Just, I., Gerhard, R., 2011. Cytotoxic properties of C. difficile TcdA lacking the proposed receptor binding domain. Naunyn-Schmiedebergs Archives of Pharmacology 383, 89-89.

Olling, A., Seehase, S., Minton, N.P., Tatge, H., Schroter, S., Kohlscheen, S., Pich, A., Just, I., Gerhard, R., 2012. Release of TcdA and TcdB from Clostridium difficile cdi 630 is not affected by functional inactivation of the tcdE gene. Microbial Pathogenesis 52, 92-100.

Olmedo, G., Ninfa, E.G., Stock, J., Youngman, P., 1990. Novel Mutations That Alter the Regulation of Sporulation in Bacillus subtilis - Evidence That Phosphorylation of Regulatory Protein Spooa Controls the Initiation of Sporulation. J Mol Biol 215, 359-372.

Osgood, D.P., Wood, N.P., Sperry, J.F., 1993. Nutritional Aspects of Cytotoxin Production by Clostridium difficile. Appl Environ Microbiol 59, 3985-3988.

Owens, R.C., Jr., Donskey, C.J., Gaynes, R.P., Loo, V.G., Muto, C.A., 2008. Antimicrobial-associated risk factors for Clostridium difficile infection. Clin Infect Dis 46 Suppl 1, S19-31.

Panis, Y., Hautefeuille, P., Hecht, Y., Lehouelleur, J., Gompel, H., 1992. Surgical-Treatment for Toxic Megacolon Complicating Pseudomembranous Colitis - Case-Report and Review of the Literature. Annales De Chirurgie 46, 453-457.

Papatheodorou, P., Carette, J.E., Bell, G.W., Schwan, C., Guttenberg, G., Brummelkamp, T.R., Aktories, K., 2011. Lipolysis-stimulated lipoprotein receptor (LSR) is the host receptor for the binary toxin Clostridium difficile transferase (CDT). Proc Natl Acad Sci U S A 108, 16422-16427.

Papatheodorou, P., Zamboglou, C., Genisyuerek, S., Guttenberg, G., Aktories, K., 2010. Clostridial Glucosylating Toxins Enter Cells via Clathrin-Mediated Endocytosis. Plos One 5.

Patel, N.C., Griesbach, C.L., DiBaise, J.K., Orenstein, R., 2013. Fecal Microbiota Transplant for Recurrent Clostridium difficile Infection: Mayo Clinic in Arizona Experience. Mayo Clinic Proceedings 88, 799-805.

Peltier, J., Courtin, P., El Meouche, I., Lemee, L., Chapot-Chartier, M.P., Pons, J.L., 2011. Clostridium difficile Has an Original Peptidoglycan Structure with a High Level of N-Acetylglucosamine Deacetylation and Mainly 3-3 Cross-links. Journal of Biological Chemistry 286, 29053-29062.

Perego, M., Spiegelman, G.B., Hoch, J.A., 1988. Structure of the Gene for the Transition-State Regulator, Abrb - Regulator Synthesis Is Controlled by the Spooa Sporulation Gene in Bacillus subtilis. Mol Microbiol 2, 689-699.

Perelle, S., Gibert, M., Bourlioux, P., Corthier, G., Popoff, M.R., 1997. Production of a complete binary toxin (actin-specific ADP-ribosyltransferase) by Clostridium difficile CD196. Infect Immun 65, 1402-1407.

184

Pettit, L.J., Browne, H.P., Yu, L., Smits, W.K., Fagan, R.P., Barquist, L., Martin, M.J., Goulding, D., Duncan, S.H., Flint, H.J., Dougan, G., Choudhary, J.S., Lawley, T.D., 2014. Functional genomics reveals that Clostridium difficile Spo0A coordinates sporulation, virulence and metabolism. Bmc Genomics 15, 160.

Planche, T., Wilcox, M.H., 2015. Diagnostic Pitfalls in Clostridium difficile Infection. Infectious Disease Clinics of North America 29, 63-+.

Pogliano, K., Harry, E., Losick, R., 1995. Visualization of the subcellular location of sporulation proteins in Bacillus subtilis using immunofluorescence microscopy. Mol Microbiol 18, 459-470.

Popoff, M.R., Rubin, E.J., Gill, D.M., Boquet, P., 1988. Actin-specific ADP-ribosyltransferase produced by a Clostridium difficile strain. Infect Immun 56, 2299-2306.

Pothoulakis, C., Lamont, J.T., 2001. Microbes and microbial toxins: paradigms for microbial-mucosal interactions II. The integrated response of the intestine to Clostridium difficile toxins. Am J Physiol Gastrointest Liver Physiol 280, G178-183.

Poxton, I.R., McCoubrey, J., Blair, G., 2001. The pathogenicity of Clostridium difficile. Clin Microbiol Infect 7, 421-427.

Punta, M., Coggill, P.C., Eberhardt, R.Y., Mistry, J., Tate, J., Boursnell, C., Pang, N., Forslund, K., Ceric, G., Clements, J., Heger, A., Holm, L., Sonnhammer, E.L., Eddy, S.R., Bateman, A., Finn, R.D., 2012. The Pfam protein families database. Nucleic Acids Res 40, D290-301.

Purcell, E.B., Mckee, R.W., McBride, S.M., Waters, C.M., Tamayo, R., 2012. Cyclic Diguanylate Inversely Regulates Motility and Aggregation in Clostridium difficile. J Bacteriol 194, 3307-3316.

Purdy, D., O'Keeffe, T.A., Elmore, M., Herbert, M., McLeod, A., Bokori-Brown, M., Ostrowski, A., Minton, N.P., 2002. Conjugative transfer of clostridial shuttle vectors from Escherichia coli to Clostridium difficile through circumvention of the restriction barrier. Mol Microbiol 46, 439-452.

Qa'Dan, M., Spyres, L.M., Ballard, J.D., 2000. pH-induced conformational changes in Clostridium difficile toxin B. Infect Immun 68, 2470-2474.

Quera, R., Espinoza, R., Estay, C., Rivera, D., 2014. Bacteremia as an adverse event of fecal microbiota transplantation in a patient with Crohn's disease and recurrent Clostridium difficile infection. J Crohns Colitis 8, 252-253.

Ransom, E.M., Ellermeier, C.D., Weiss, D.S., 2015. Use of mCherry Red fluorescent protein for studies of protein localization and gene expression in Clostridium difficile. Appl Environ Microbiol 81, 1652-1660.

Ransom, E.M., Williams, K.B., Weiss, D.S., Ellermeier, C.D., 2014. Identification and characterization of a gene cluster required for proper rod shape, cell division, and pathogenesis in Clostridium difficile. J Bacteriol 196, 2290-2300.

Ratnayake-Lecamwasam, M., Serror, P., Wong, K.W., Sonenshein, A.L., 2001. Bacillus subtilis CodY represses early-stationary-phase genes by sensing GTP levels. Genes Dev 15, 1093-1103.

Rausch, S., Kaibie, T., 2014. Clostridium difficile-associated, toxic Megacolon after Cystoprostatectomy and Attachment of Ileum-Conduits. Aktuelle Urologie 45, 506-507.

185

Ravagnani, A., Jennert, K.C.B., Steiner, E., Grunberg, R., Jefferies, J.R., Wilkinson, S.R., Young, D.I., Tidswell, E.C., Brown, D.P., Youngman, P., Morris, J.G., Young, M., 2000. Spo0A directly controls the switch from acid to solvent production in solvent-forming clostridia. Mol Microbiol 37, 1172-1185.

Reddy, M., 2007. Role of FtsEX in cell division of Escherichia coli: viability of ftsEX mutants is dependent on functional SufI or high osmotic strength. J Bacteriol 189, 98-108.

Ricard, M., Hirota, Y., 1973. Process of cellular division in Escherichia coli: physiological study on thermosensitive mutants defective in cell division. J Bacteriol 116, 314-322.

Riggs, M.M., Sethi, A.K., Zabarsky, T.F., Eckstein, E.C., Jump, R.L.P., Donskey, C.J., 2007. Asymptomatic carriers are a potential source for transmission of epidemic and nonepidemic Clostridium difficile strains among long-term care facility residents. Clinical Infectious Diseases 45, 992-998.

Roach, E.J., Kimber, M.S., Khursigara, C.M., 2014. Crystal structure and site-directed mutational analysis reveals key residues involved in Escherichia coli ZapA function. J Biol Chem 289, 23276-23286.

Rohlke, F., Stollman, N., 2012. Fecal microbiota transplantation in relapsing Clostridium difficile infection. Therapeutic Advances in Gastroenterology 5, 403-420.

Rosenbusch, K.E., Bakker, D., Kuijper, E.J., Smits, W.K., 2012. C. difficile 630Deltaerm Spo0A regulates sporulation, but does not contribute to toxin production, by direct high-affinity binding to target DNA. Plos One 7, e48608.

Rueda, S., Vicente, M., Mingorance, J., 2003. Concentration and assembly of the division ring proteins FtsZ, FtsA, and ZipA during the Escherichia coli cell cycle. J Bacteriol 185, 3344-3351.

Ruiz, C.G., Lopez, S.E.R., Villella, F.V., Carballal, C.M., Mori, R.V., Juarez, M.C.D., Fustegueras, J.A.B., Kissler, J.J.O., 2015. Fatal pseudomembranous colitis in immunocompetent patient. Revista Espanola De Enfermedades Digestivas 107, 117-119.

Rupnik, M., Pabst, S., Rupnik, M., von Eichel-Streiber, C., Urlaub, H., Soling, H.D., 2005. Characterization of the cleavage site and function of resulting cleavage fragments after limited proteolysis of Clostridium difficile toxin B (TcdB) by host cells. Microbiology-Sgm 151, 199-208.

Rupnik, M., Wilcox, M.H., Gerding, D.N., 2009. Clostridium difficile infection: new developments in epidemiology and pathogenesis. Nat Rev Microbiol 7, 526-536.

Saile, E., Koehler, T.M., 2002. Control of anthrax toxin gene expression by the transition state regulator abrB. J Bacteriol 184, 370-380.

Sambol, S.P., Tang, J.K., Merrigan, M.M., Johnson, S., Gerding, D.N., 2001. Infection of hamsters with epidemiologically important strains of Clostridium difficile. J Infect Dis 183, 1760-1766.

Sastalla, I., Chim, K., Cheung, G.Y., Pomerantsev, A.P., Leppla, S.H., 2009. Codon-optimized fluorescent proteins designed for expression in low-GC gram-positive bacteria. Appl Environ Microbiol 75, 2099-2110.

Sauerborn, M., Leukel, P., von Eichel-Streiber, C., 1997. The C-terminal ligand-binding domain of Clostridium difficile toxin A (TcdA) abrogates TcdA-specific binding to cells and prevents mouse lethality. FEMS Microbiol Lett 155, 45-54.

186

Saujet, L., Monot, M., Dupuy, B., Soutourina, O., Martin-Verstraete, I., 2011. The key sigma factor of transition phase, SigH, controls sporulation, metabolism, and virulence factor expression in Clostridium difficile. J Bacteriol 193, 3186-3196.

Savidge, T.C., Pan, W.H., Newman, P., O'Brien, M., Anton, P.M., Pothoulakis, C., 2003. Clostridium difficile toxin B is an inflammatory enterotoxin in human intestine. Gastroenterology 125, 413-420.

Schaffer, J., Olliver, P., Keck, G., 1987. Studies in Infants and Children with Clostridium difficile Toxin in Stools. Microecology and Therapy, Vol 17 17, 57-57.

Scheffers, D.J., 2008. The effect of MinC on FtsZ polymerization is pH dependent and can be counteracted by ZapA. FEBS Lett 582, 2601-2608.

Schumacher, M.A., Allen, G.S., Diel, M., Seidel, G., Hillen, W., Brennan, R.G., 2004. Structural basis for allosteric control of the transcription regulator CcpA by the phosphoprotein HPr-Ser46-P. Cell 118, 731-741.

Schuster-Bockler, B., Schultz, J., Rahmann, S., 2004. HMM Logos for visualization of protein families. Bmc Bioinformatics 5, 7.

Sebaihia, M., Wren, B.W., Mullany, P., Fairweather, N.F., Minton, N., Stabler, R., Thomson, N.R., Roberts, A.P., Cerdeno-Tarraga, A.M., Wang, H., Holden, M.T., Wright, A., Churcher, C., Quail, M.A., Baker, S., Bason, N., Brooks, K., Chillingworth, T., Cronin, A., Davis, P., Dowd, L., Fraser, A., Feltwell, T., Hance, Z., Holroyd, S., Jagels, K., Moule, S., Mungall, K., Price, C., Rabbinowitsch, E., Sharp, S., Simmonds, M., Stevens, K., Unwin, L., Whithead, S., Dupuy, B., Dougan, G., Barrell, B., Parkhill, J., 2006. The multidrug-resistant human pathogen Clostridium difficile has a highly mobile, mosaic genome. Nat Genet 38, 779-786.

Sham, L.T., Barendt, S.M., Kopecky, K.E., Winkler, M.E., 2011. Essential PcsB putative peptidoglycan hydrolase interacts with the essential FtsXSpn cell division protein in Streptococcus pneumoniae D39. Proc Natl Acad Sci U S A 108, E1061-1069.

Shaner, N.C., Campbell, R.E., Steinbach, P.A., Giepmans, B.N., Palmer, A.E., Tsien, R.Y., 2004. Improved monomeric red, orange and yellow fluorescent proteins derived from Discosoma sp. red fluorescent protein. Nat Biotechnol 22, 1567-1572.

Shaner, N.C., Steinbach, P.A., Tsien, R.Y., 2005. A guide to choosing fluorescent proteins. Nat Methods 2, 905-909.

Shcherbo, D., Murphy, C.S., Ermakova, G.V., Solovieva, E.A., Chepurnykh, T.V., Shcheglov, A.S., Verkhusha, V.V., Pletnev, V.Z., Hazelwood, K.L., Roche, P.M., Lukyanov, S., Zaraisky, A.G., Davidson, M.W., Chudakov, D.M., 2009. Far-red fluorescent tags for protein imaging in living tissues. Biochemical Journal 418, 567-574.

Sickmier, E.A., Brekasis, D., Paranawithana, S., Bonanno, J.B., Paget, M.S., Burley, S.K., Kielkopf, C.L., 2005. X-ray structure of a Rex-family repressor/NADH complex insights into the mechanism of redox sensing. Structure 13, 43-54.

Siegrist, M.S., Swarts, B.M., Fox, D.M., Lim, S.A., Bertozzi, C.R., 2015. Illumination of growth, division and secretion by metabolic labeling of the bacterial cell surface. Fems Microbiology Reviews 39, 184-202.

187

Siegrist, M.S., Whiteside, S., Jewett, J.C., Aditham, A., Cava, F., Bertozzi, C.R., 2013. D-Amino Acid Chemical Reporters Reveal Peptidoglycan Dynamics of an Intracellular Pathogen. Acs Chemical Biology 8, 500-505.

Sievers, F., Wilm, A., Dineen, D., Gibson, T.J., Karplus, K., Li, W., Lopez, R., McWilliam, H., Remmert, M., Soding, J., Thompson, J.D., Higgins, D.G., 2011. Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal Omega. Mol Syst Biol 7, 539.

Small, E., Marrington, R., Rodger, A., Scott, D.J., Sloan, K., Roper, D., Dafforn, T.R., Addinall, S.G., 2007. FtsZ polymer-bundling by the Escherichia coli ZapA orthologue, YgfE, involves a conformational change in bound GTP. J Mol Biol 369, 210-221.

Smith, R.M., O'Hare, J.P., Cope, O., Wesselhoeft, C., Alt, R.E., 1953. ANTIBIOTICS, Staphylococcal enteritis and pseudomembranous enterocolitis. N Engl J Med 249, 37-40.

Snyder, M.L., 1937. Further studies on Bacillus difficilis (Hall and O'Toole). Journal of Infectious Diseases 60, 223-231.

Spigaglia, P., Barketi-Klai, A., Collignon, A., Mastrantonio, P., Barbanti, F., Rupnik, M., Janezic, S., Kansau, I., 2013. Surface-layer (S-layer) of human and animal Clostridium difficile strains and their behaviour in adherence to epithelial cells and intestinal colonization. J Med Microbiol 62, 1386-1393.

Stabler, R.A., He, M., Dawson, L., Martin, M., Valiente, E., Corton, C., Lawley, T.D., Sebaihia, M., Quail, M.A., Rose, G., Gerding, D.N., Gibert, M., Popoff, M.R., Parkhill, J., Dougan, G., Wren, B.W., 2009. Comparative genome and phenotypic analysis of Clostridium difficile 027 strains provides insight into the evolution of a hypervirulent bacterium. Genome Biology 10.

Starnbach, M.N., Lory, S., 1992. The Flia (Rpof) Gene of Pseudomonas aeruginosa Encodes an Alternative Sigma Factor Required for Flagellin Synthesis. Mol Microbiol 6, 459-469.

Steele, J., Mukherjee, J., Parry, N., Tzipori, S., 2013. Antibody Against TcdB, but Not TcdA, Prevents Development of Gastrointestinal and Systemic Clostridium difficile Disease. Journal of Infectious Diseases 207, 323-330.

Stewart, P.S., Franklin, M.J., 2008. Physiological heterogeneity in biofilms. Nature Reviews Microbiology 6, 199-210.

Stollman, N., Surawicz, C., 2012. Fecal Transplant for Clostridium difficile. Archives of Internal Medicine 172, 825-825.

Strahl, H., Hamoen, L.W., 2010. Membrane potential is important for bacterial cell division. Proc Natl Acad Sci U S A 107, 12281-12286.

Stubbs, S., Rupnik, M., Gibert, M., Brazier, J., Duerden, B., Popoff, M., 2000. Production of actin-specific ADP-ribosyltransferase (binary toxin) by strains of Clostridium difficile. FEMS Microbiol Lett 186, 307-312.

Tal, S., Gurevich, A., Guller, V., Gurevich, I., Berger, D., Levi, S., 2002. Risk factors for recurrence of Clostridium difficile-associated diarrhea in the elderly. Scandinavian Journal of Infectious Diseases 34, 594-597.

Tan, K.S., Wee, B.Y., Song, K.P., 2001. Evidence for holin function of tcdE gene in the pathogenicity of Clostridium difficile. J Med Microbiol 50, 613-619.

188

Tarry, M., Arends, S.J., Roversi, P., Piette, E., Sargent, F., Berks, B.C., Weiss, D.S., Lea, S.M., 2009. The Escherichia coli cell division protein and model Tat substrate SufI (FtsP) localizes to the septal ring and has a multicopper oxidase-like structure. J Mol Biol 386, 504-519.

Tasteyre, A., Barc, M.C., Collignon, A., Boureau, H., Karjalainen, T., 2001. Role of FliC and FliD flagellar proteins of Clostridium difficile in adherence and gut colonization. Infect Immun 69, 7937-7940.

Tasteyre, A., Karjalainen, T., Avesani, V., Delmee, M., Collignon, A., Bourlioux, P., Barc, M.C., 2000. Phenotypic and genotypic diversity of the flagellin gene (fliC) among Clostridium difficile isolates from different serogroups. Journal of Clinical Microbiology 38, 3179-3186.

Tleyjeh, I.M., Bin Abdulhak, A.A., Riaz, M., Alasmari, F.A., Garbati, M.A., AlGhamdi, M., Khan, A.R., Al Tannir, M., Erwin, P.J., Ibrahim, T., AlLehibi, A., Baddour, L.M., Sutton, A.J., 2012. Association between Proton Pump Inhibitor Therapy and Clostridium difficile Infection: A Contemporary Systematic Review and Meta-Analysis. Plos One 7.

Treloar, A.J., Kalra, L., 1987. Mortality and Clostridium difficile Diarrhea in the Elderly. Lancet 2, 1279-1279.

Triadafilopoulos, G., Pothoulakis, C., O'Brien, M.J., LaMont, J.T., 1987. Differential effects of Clostridium difficile toxins A and B on rabbit ileum. Gastroenterology 93, 273-279.

Trieu-Cuot, P., Arthur, M., Courvalin, P., 1987. Origin, evolution and dissemination of antibiotic resistance genes. Microbiol Sci 4, 263-266.

Tsien, R.Y., 1998. The green fluorescent protein. Annu Rev Biochem 67, 509-544.

Tullus, K., Aronsson, B., Marcus, S., Mollby, R., 1989. Intestinal Colonization with Clostridium difficile in Infants up to 18 Months of Age. European Journal of Clinical Microbiology & Infectious Diseases 8, 390-393.

Twine, S.M., Reid, C.W., Aubry, A., McMullin, D.R., Fulton, K.M., Austin, J., Logan, S.M., 2009. Motility and Flagellar Glycosylation in Clostridium difficile. J Bacteriol 191, 7050-7062.

Uehara, T., Parzych, K.R., Dinh, T., Bernhardt, T.G., 2010. Daughter cell separation is controlled by cytokinetic ring-activated cell wall hydrolysis. Embo Journal 29, 1412-1422.

Uliczka, F., Pisano, F., Kochut, A., Opitz, W., Herbst, K., Stolz, T., Dersch, P., 2011. Monitoring of Gene Expression in Bacteria during Infections Using an Adaptable Set of Bioluminescent, Fluorescent and Colorigenic Fusion Vectors. Plos One 6.

Ursinus, A., van den Ent, F., Brechtel, S., de Pedro, M., Höltje, J.V., Löwe, J., Vollmer, W., 2004. Murein (peptidoglycan) binding property of the essential cell division protein FtsN from Escherichia coli. J Bacteriol 186, 6728-6737.

van Eijk, E., Anvar, S.Y., Browne, H.P., Leung, W.Y., Frank, J., Schmitz, A.M., Roberts, A.P., Smits, W.K., 2015. Complete genome sequence of the Clostridium difficile laboratory strain 630 Delta erm reveals differences from strain 630, including translocation of the mobile element CTn5. Bmc Genomics 16.

van Nood, E., Vrieze, A., Nieuwdorp, M., Fuentes, S., Zoetendal, E.G., de Vos, W.M., Visser, C.E., Kuijper, E.J., Bartelsman, J.F., Tijssen, J.G., Speelman, P., Dijkgraaf, M.G., Keller, J.J., 2013. Duodenal infusion of donor feces for recurrent Clostridium difficile. N Engl J Med 368, 407-415.

189

van Schaik, W., Chateau, A., Dillies, M.A., Coppee, J.Y., Sonenshein, A.L., Fouet, A., 2009. The Global Regulator CodY Regulates Toxin Gene Expression in Bacillus anthracis and Is Required for Full Virulence. Infect Immun 77, 4437-4445.

Veening, J.W., Hamoen, L.W., Kuipers, O.P., 2005. Phosphatases modulate the bistable sporulation gene expression pattern in Bacillus subtilis. Mol Microbiol 56, 1481-1494.

Velanovich, V., Laporta, A.J., Garrett, W.L., Richards, T.B., Cornett, P.A., 1992. Pseudomembranous Colitis Leading to Toxic Megacolon Associated with Antineoplastic Chemotherapy - Report of a Case and Review of the Literature. Diseases of the Colon & Rectum 35, 369-372.

Vellanoweth, R.L., Rabinowitz, J.C., 1992. The influence of ribosome-binding-site elements on translational efficiency in Bacillus subtilis and Escherichia coli in vivo. Mol Microbiol 6, 1105-1114.

von Eichel-Streiber, C., Laufenberg-Feldmann, R., Sartingen, S., Schulze, J., Sauerborn, M., 1992. Comparative sequence analysis of the Clostridium difficile toxins A and B. Mol Gen Genet 233, 260-268.

Voth, D.E., Ballard, J.D., 2005. Clostridium difficile toxins: Mechanism of action and role in disease. Clinical Microbiology Reviews 18, 247-+.

Walker, A.S., Eyre, D.W., Wyllie, D.H., Dingle, K.E., Griffiths, D., Shine, B., Oakley, S., O'Connor, L., Finney, J., Vaughan, A., Crook, D.W., Wilcox, M.H., Peto, T.E.A., IO, I.O.R.D., 2013. Relationship Between Bacterial Strain Type, Host Biomarkers, and Mortality in Clostridium difficile Infection. Clinical Infectious Diseases 56, 1589-1600.

Warny, M., Keates, A.C., Pothoulakis, C., LaMont, J.T., Kelly, C.P., 1997. Rho glucosylation by Clostridium difficile toxins A and B triggers CD14 shedding by human monocytes. Gastroenterology 112, A1115-A1115.

Warny, M., Pepin, J., Fang, A., Killgore, G., Thompson, A., Brazier, J., Frost, E., McDonald, L.C., 2005. Toxin production by an emerging strain of Clostridium difficile associated with outbreaks of severe disease in North America and Europe. Lancet 366, 1079-1084.

Weil, A.A., Hohmann, E.L., 2015. Fecal Microbiota Transplant: Benefits and Risks. Open Forum Infectious Diseases 2.

Weir, J., Predich, M., Dubnau, E., Nair, G., Smith, I., 1991. Regulation of Spo0h, a Gene Coding for the Bacillus subtilis Sigma-H Factor. J Bacteriol 173, 521-529.

Weiss, D.S., Chen, J.C., Ghigo, J.M., Boyd, D., Beckwith, J., 1999. Localization of FtsI (PBP3) to the septal ring requires its membrane anchor, the Z ring, FtsA, FtsQ, and FtsL. J Bacteriol 181, 508-520.

Wilcox, M.H., Shetty, N., Fawley, W.N., Shemko, M., Coen, P., Birtles, A., Cairns, M., Curran, M.D., Dodgson, K.J., Green, S.M., Hardy, K.J., Hawkey, P.M., Magee, J.G., Sails, A.D., Wren, M.W.D., 2012. Changing Epidemiology of Clostridium difficile Infection Following the Introduction of a National Ribotyping-Based Surveillance Scheme in England. Clinical Infectious Diseases 55, 1056-1063.

Wilkins, T.D., Tucker, K.D., 1989. Clostridium difficile Toxin-a (Enterotoxin) Uses Gal-Alpha-1-3gal-Beta-1-4g1cnac as a Functional Receptor. Microecology and Therapy, Vol 19 19, 225-227.

190

Williams, K.B., Yahashiri, A., Arends, S.J., Popham, D.L., Fowler, C.A., Weiss, D.S., 2013. Nuclear magnetic resonance solution structure of the peptidoglycan-binding SPOR domain from Escherichia coli DamX: insights into septal localization. Biochemistry 52, 627-639.

Wilson, K.H., Kennedy, M.J., Fekety, F.R., 1982. Use of Sodium Taurocholate to Enhance Spore Recovery on a Medium Selective for Clostridium difficile. Journal of Clinical Microbiology 15, 443-446.

Wilson, M.J., Carlson, P.E., Janes, B.K., Hanna, P.C., 2012. Membrane Topology of the Bacillus anthracis GerH Germinant Receptor Proteins. J Bacteriol 194, 1369-1377.

Winn, W.C., Koneman, E.W., 2006. Koneman's color atlas and textbook of diagnostic microbiology, 6th ed. Lippincott Williams & Wilkins, Philadelphia.

Wohlan, K., Goy, S., Olling, A., Srivaratharajan, S., Tatge, H., Genth, H., Gerhard, R., 2014. Pyknotic cell death induced by Clostridium difficile TcdB: chromatin condensation and nuclear blister are induced independently of the glucosyltransferase activity. Cellular Microbiology 16, 1678-1692.

Wullt, M., Odenholt, I., Walder, M., 2003. Activity of three disinfectants and acidified nitrite against Clostridium difficile spores. Infection Control and Hospital Epidemiology 24, 765-768.

Yang, D.C., Peters, N.T., Parzych, K.R., Uehara, T., Markovski, M., Bernhardt, T.G., 2011. An ATP-binding cassette transporter-like complex governs cell-wall hydrolysis at the bacterial cytokinetic ring. Proc Natl Acad Sci U S A 108, E1052-1060.

Yearsley, K.A., Gilby, L.J., Ramadas, A.V., Kubiak, E.M., Fone, D.L., Allison, M.C., 2006. Proton pump inhibitor therapy is a risk factor for Clostridium difficile-associated diarrhoea. Alimentary Pharmacology & Therapeutics 24, 613-619.

Yu, X.C., Margolin, W., 1999. FtsZ ring clusters in min and partition mutants: role of both the Min system and the nucleoid in regulating FtsZ ring localization. Mol Microbiol 32, 315-326.

Yuan, P.F., Zhang, H.M., Cai, C.Z., Zhu, S.Y., Zhou, Y.X., Yang, X.Z., He, R.N., Li, C., Guo, S.J., Li, S., Huang, T.X., Perez-Cordon, G., Feng, H.P., Wei, W.S., 2015. Chondroitin sulfate proteoglycan 4 functions as the cellular receptor for Clostridium difficile toxin B. Cell Research 25, 157-168.

Yutin, N., Galperin, M.Y., 2013. A genomic update on clostridial phylogeny: Gram-negative spore formers and other misplaced clostridia. Environ Microbiol 15, 2631-2641.

Zhang, C., Xing, X.H., Lou, K., 2005. Rapid detection of a gfp-marked Enterobacter aerogenes under anaerobic conditions by aerobic fluorescence recovery. FEMS Microbiol Lett 249, 211-218.

Zidaric, V., Pardon, B., dos Vultos, T., Deprez, P., Brouwer, M.S.M., Roberts, A.P., Henriques, A.O., Rupnik, M., 2012. Different Antibiotic Resistance and Sporulation Properties within Multiclonal Clostridium difficile PCR Ribotypes 078, 126, and 033 in a Single Calf Farm. Appl Environ Microbiol 78, 8515-8522.

Zidaric, V., Zemljic, M., Janezic, S., Kocuvan, A., Rupnik, M., 2008. High diversity of Clostridium difficile genotypes isolated from a single poultry farm producing replacement laying hens. Anaerobe 14, 325-327.