Isabelle Martin

Anna Audrey

Claire

Marc Olga Julian

Transito Ritsuko

Miguel Teixeira

Marisa Flávia

Marta Sofia

Jorge Pereira

Jorge Filipe

Alicia André

Rbr Fecys

Fe -Fe Rbr

Fe -Fe Rd

Dfx Fpra

Rbr

Spo0a Rsta

Cotb Cota

Cdeb Cdea

Bcla1

Reduction Dismutation SOD

Keywords: Clostridium difficile, Manteau de la spore, Morphogénèse, CotL, Oxygène, ERO, détoxification, Rubrerythrines, protéine Flavodiiron Dysbiosis Stomach Small intestine Duodenum Ileum Large intestine Cecum Colon Rectum Dormant spores Germinating spores Vegetative cells

Clostridium difficile qui est une bactérie anaérobie stricte et sporulante, est la principale cause des diarrhées associées à la prise d'antibiotiques. Les spores jouent un rôle central dans le cycle infectieux de C. difficile : résistance en dehors de l'hôte, persistance des bactéries et transmission de l'infection. Lorsqu'elles sont ingérées par l'hôte, les spores de C. difficile germent dans le côlon en présence de certains sels biliaires pour former des cellules végétatives qui vont produire des toxines responsables des symptômes.

Les cellules végétatives sont exposées à différents stress incluant les espèces réactives de l'oxygène et de l'azote (ROS/RNS) produites par le système immunitaire de l'hôte lors de l'inflammation. De plus, bien que le tractus intestinal soit considéré comme anoxique, une faible teneur en oxygène (O 2 ) est présente dans l'intestin et elle va augmenter après un traitement antibiotique. Pour survivre dans cet environnement oxydant hostile, C. difficile a développé des mécanismes de protection, des voies de détoxification et des systèmes de réparation. Le facteur sigma alternatif de la réponse générale au stress, σ B , joue un rôle important dans la protection de C. difficile face aux stress rencontrés dans l'hôte incluant l'O 2 et les ROS. Parmi les gènes contrôlés positivement par σ B , nous avons identifié des gènes codant pour des protéines probablement impliquées dans la tolérance à l'O 2 et la détoxification des ROS, comme les deux réverses rubrérythrines (revRbrs), CD1524 et CD1474, et les protéines de type flavodiiron (FDPs), CD1157 et CD1623. Nous avons montré que les gènes codant les deux revRbrs et CD1623 sont exprimés sous le contrôle de σ B et de manière hétérogène dans la population bactérienne. Le gène CD1157 est lui exprimé à partir de deux promoteurs dépendants l'un de σ A et l'autre de σ B . Nous avons ensuite montré que CD1157 a une activité O 2 -réductase alors que les deux revRbrs ont une activité H 2 O 2 -et O 2réductase, l'H 2 O 2 étant le substrat préféré. De plus, nous avons montré qu'un double mutant ∆CD1474-∆CD1524 et un mutant CD1157 sont moins tolérants à une faible tension en O 2 et que le mutant ∆CD1623 est plus sensible à l'exposition à l'air. Ces résultats démontrent le rôle clé de ces protéines dans la tolérance à l'O 2 et la réponse au stress oxydant chez C. difficile.

Au cours du cycle infectieux, dans le côlon, certaines cellules végétatives sporulent. Les couches de surface des spores, le manteau et l'exosporium, permettent aux spores de résister aux stress physiques et chimiques. Cependant, peu de choses sont connues sur leurs mécanismes d'assemblage. Dans ce travail, nous avons caractérisé une nouvelle protéine, CotL, qui est nécessaire pour l'assemblage des couches externes de la spore. Nous avons montré que le gène cotL est exprimé dans la cellule mère sous le double contrôle des facteurs sigma de sporulation, σ E et σ K . La protéine CotL est localisée dans le manteau de la spore, et les spores du mutant cotL ont un défaut morphologique majeur au niveau des couches du manteau et de l'exosporium. De plus, les spores du mutant contiennent une quantité réduite de plusieurs protéines du manteau et de l'exosporium et présentent un défaut dans leur localisation dans des cellules en cours de sporulation. Enfin, les spores du mutant cotL sont plus sensibles au lysozyme et présentent un défaut de germination, un phénotype vraisemblablement associé à la modification de la structure des couches externes de la spore. Ces résultats suggèrent fortement que CotL est une protéine morphogénétique essentielle à l'assemblage des couches externes de la spore chez C. difficile.

Stephens 1998 A B

List of abbreviations

Table of Contents

Chapter 1: The enteric pathogen Clostridium difficile

The human body contains a vast array of microorganisms but the highest density and diversity of microbial species inhabiting us is present in the intestine (10 12-14 bacteria/gr in the colon). A typical human intestinal microbiota contains up to 1000 species of commensal, beneficial and pathogenic organisms [START_REF] Qin | A human gut microbial gene catalogue established by metagenomic sequencing[END_REF]. Most of these microorganisms belong to only two phyla: the Bacteroidetes (genus Bacteroides) and the Firmicutes (genus Clostridium, Faecalibacterium, Lactobacillus, Ruminococcus). Actinobacteria and Proteobacteria are also represented, but as minor constituents [START_REF] Lozupone | Diversity, stability and resilience of the human gut microbiota[END_REF]. A health-associated intestinal microbiota form a community that helps in food digestion, protects against enteropathogens and stimulate the immune system [START_REF] Backhed | The gut microbiota as an environmental factor that regulates fat storage[END_REF][START_REF] Littman | Role of the commensal microbiota in normal and pathogenic host immune responses[END_REF][START_REF] Stecher | Mechanisms controlling pathogen colonization of the gut[END_REF]. The host, in turn, provides habitat and nutrients. Recent studies suggest that at least half of the bacterial genera found in the gut produce spores [START_REF] Browne | Culturing of 'unculturable' human microbiota reveals novel taxa and extensive sporulation[END_REF]. Most of these spore formers present in gut correspond to clostridial species including Clostridium difficile.

I. Clostridium difficile

C. difficile is a gram-positive, toxin-producing and spore-forming anaerobe. C. difficile was identified and described for the first time in 1935 by Hall and O'Toole as part of the intestinal microflora in neonates [START_REF] Hall | Intestinal flora in new-born infants with a description of a new pathogenic anaerobic, Bacillus diffıcilis[END_REF]. At that time, the bacterium was described as an "actively motile, heavy-bodied rod with elongated subterminal or nearly terminal spores" (Figure 1A). This specie was named Bacillus difficilis to reflect the difficulties in its isolation and growth in the laboratory. C. difficile bacillus is between 3 and 15 µm long and 0.5 to 2 µm wide and colonies are opaque and irregular in Petri dish (Figure 1B). In 1970's, multiple studies provided evidence that C. difficile was the etiologic agent of antibiotic-associated diarrhea and colitis not only in animals, but also in humans [START_REF] Carroll | Biology of Clostridium difficile: implications for epidemiology and diagnosis[END_REF].

Nowadays, C. difficile is recognized as the leading cause of nosocomial antibiotic-associated diarrhea in developed countries [START_REF] Smits | Clostridium difficile infection[END_REF]. The development of C. difficile infection (CDI) is associated with different virulence factors, mainly with the release of two toxins, TcdA and TcdB, that play a crucial role in the pathogenicity of C. difficile [START_REF] Voth | Clostridium difficile toxins: mechanism of action and role in disease[END_REF][START_REF] Aktories | Clostridium difficile Toxin Biology[END_REF][START_REF] Chandrasekaran | The role of toxins in Clostridium difficile infection[END_REF].

C. difficile belongs to the cluster XI of clostridia with, among others, Clostridium sticklandii and Clostridium sordelii [START_REF] Collins | The phylogeny of the genus Clostridium: proposal of five new genera and eleven new species combinations[END_REF]. In 2009, C. difficile and the related Cluster XI were reclassified into a family-level group, Peptostreptococcaceae, distinct from the current Clostridiaceae family. Later on, it was renamed Peptoclostridium difficile [START_REF] Yutin | A genomic update on clostridial phylogeny: Gram-negative spore formers and other misplaced clostridia[END_REF]. More recently, C. difficile was renamed Clostridioides difficile and is considered the type species of the genus Clostridioides [START_REF] Lawson | Reclassification of Clostridium difficile as Clostridioides difficile (Hall and O'Toole 1935) Prevot[END_REF]. For clarity in the manuscript, I will use the more familiar designation Clostridium difficile and C. difficile. The complete genome of C. difficile reference 630 strain was sequenced in 2006 [START_REF] Sebaihia | The multidrug-resistant human pathogen Clostridium difficile has a highly mobile, mosaic genome[END_REF]). The chromosome is composed by 4 290 252 base pairs (bp) with a G+C content of 29%, and 3 776 coding sequences are present [START_REF] Monot | Reannotation of the genome sequence of Clostridium difficile strain 630[END_REF][START_REF] Dannheim | Manual curation and reannotation of the genomes of Clostridium difficile 630Deltaerm and C. difficile 630[END_REF]. C.

difficile shares only 15% of its coding sequences with Clostridium botulinum and Clostridium tetani, and 50% of the coding sequences are unique to this bacterium. Moreover, 11% of the chromosome consists of mobile genetic elements [START_REF] Sebaihia | The multidrug-resistant human pathogen Clostridium difficile has a highly mobile, mosaic genome[END_REF]). These mobile elements are responsible for the acquisition of genes that probably favors a better adaptation to the environment of the host's digestive tract, such as genes involved in resistance to antibacterial compounds, virulence or production of structures and surface proteins. For example, C. difficile resistance to antibiotics such as erythromycin, chloramphenicol or tetracycline is largely mediated by transposons present in the genome [START_REF] Lyras | The large resolvase TnpX is the only transposonencoded protein required for transposition of the Tn4451/3 family of integrative mobilizable elements[END_REF][START_REF] Hussain | Generation of an erythromycin-sensitive derivative of Clostridium difficile strain 630 (630Deltaerm) and demonstration that the conjugative transposon Tn916DeltaE enters the genome of this strain at multiple sites[END_REF][START_REF] Sebaihia | The multidrug-resistant human pathogen Clostridium difficile has a highly mobile, mosaic genome[END_REF].

I.1. C. difficile infection cycle

C. difficile is transmitted via the oral-faecal route (Figure 2). Being a strict anaerobe, the vegetative form of C. difficile cannot survive in an oxygenated environment for long and is then unable to transmit the infection. The infectious potential of this bacterium is linked to its ability to produce spores that play a crucial role in C. difficile transmission and persistence (Deakin et al. 2012). Spores are metabolically inactive forms, highly resistant to extremes of temperature, desiccation, oxidative stress, antibiotics, the acidic environment of the stomach as well as to disinfectants such as hypochlorite and hydrogen peroxide used to eradicate and control dissemination [START_REF] Nicholson | Resistance of Bacillus endospores to extreme terrestrial and extraterrestrial environments[END_REF], Baines et al. 2009[START_REF] Oughton | Hand hygiene with soap and water is superior to alcohol rub and antiseptic wipes for removal of Clostridium difficile[END_REF][START_REF] Carroll | Biology of Clostridium difficile: implications for epidemiology and diagnosis[END_REF][START_REF] Setlow | Spore Resistance Properties[END_REF]. C. difficile spores are widely distributed in the environment and can survive for long periods of time. They can be isolated from soil, water, food, hospital environment and the intestinal tract of both humans and domesticated animals [START_REF] Weese | Clostridium difficile in food--innocent bystander or serious threat?[END_REF].

When ingested, the spores survive the acidic pH of the stomach, and reach the intestine where IgA Bacterial metabolism they are thought to attach to the epithelial cells in order to achieve efficient colonization.

Indeed, recent studies have shown that attachment of the spore to the intestine seems to be essential for CDI development. Paredes-Sabja et al showed that the spore surface of C. difficile interacts with the surface of intestinal epithelial cells (Paredes-Sabja and Sarker 2012). More recently, it was demonstrated that the spore surface protein CotE enables binding of the spore to mucus by direct interaction with mucin [START_REF] Hong | The Spore Coat Protein CotE Facilitates Host Colonization by Clostridium difficile[END_REF]). In the intestine, when the normal gut microbiota is disturbed, as during antibiotic treatment, the ratio of the bile acids derivatives cholate (CA) and chenodeoxycholate (CDCA) increases and the spores germinate (Sorg andSonenshein 2008, Giel et al. 2010). Then vegetative cells multiply and colonize the cecum and the colon [START_REF] Koenigsknecht | Dynamics and establishment of Clostridium difficile infection in the murine gastrointestinal tract[END_REF]. C. difficile will then produce the two major virulence factors: the TcdA and TcdB toxins [START_REF] Carroll | Biology of Clostridium difficile: implications for epidemiology and diagnosis[END_REF]. These two toxins will cause damage to intestinal epithelial cells and trigger a strong host inflammatory response that is responsible for diarrhea, colitis and colon perforation (Figure 2). In the large intestine, C. difficile also produces spores during infection that will be released into the environment during diarrheal episodes allowing transmission of C. difficile to a new host.

I.2. CDI risk factors and colonization resistance against C. difficile

The life cycle of C. difficile is strongly influenced by the gut microbiota of the host and its associated metabolites [START_REF] Crobach | Understanding Clostridium difficile Colonization[END_REF][START_REF] Samarkos | The role of gut microbiota in Clostridium difficile infection[END_REF]. Individuals with a normal balanced microbiota are usually resistant to infection by C. difficile [START_REF] Carroll | Biology of Clostridium difficile: implications for epidemiology and diagnosis[END_REF]Bartlett 2011, Britton and[START_REF] Britton | Interaction between the intestinal microbiota and host in Clostridium difficile colonization resistance[END_REF]. This is because the microbial and metabolic environment of the gut is able to directly or indirectly prevent colonization by this pathogen (Figure 3) [START_REF] Britton | Interaction between the intestinal microbiota and host in Clostridium difficile colonization resistance[END_REF]. One mechanism by which the gut microbiota prevents C. difficile invasion of the intestinal tract is via the transformation of bile acids (Figure 3 and4). Humans secrete bile acids from cholesterol into the small intestine in response to consumption of meal. Two main primary bile acids are produced, CA and CDCA, that are conjugated to glycine or taurine amino acids (Figure 4). First, bile acids hydrolases produced by members of the intestinal microbiota deconjugate the bile acids from their amino acids in the intestinal lumen [START_REF] Ridlon | Bile salt biotransformations by human intestinal bacteria[END_REF]. Then, CA and CDCA are metabolized via the enzyme 7-dehydroxylase, into the secondary bile acids deoxycholate and lithocholate, respectively (Figure 4). This reaction is carried out by a small percentage of commensal bacteria such as Clostridium scindens [START_REF] Wells | Identification and characterization of a bile acid 7alphadehydroxylation operon in Clostridium sp. strain TO-931, a highly active 7alphadehydroxylating strain isolated from human feces[END_REF]Hylemon 2000, Buffie et al. 2015). In fact, a The liver synthesizes bile acids from cholesterol and secretes them as conjugated bile acids. Ileum gut microbiota produces bile salt hydrolases that deconjugate bile acids to primary bile acids, chenodeoxycholate and cholate. Primary bile acids serve as substrates for 7-dehydroxylases, made by a small subset of clostridial organisms such as C. scindens, which generate secondary bile acids lithocholate and deoxycholate. Chenodeoxycholate and secondary bile acids are toxic to vegetative cells of C. difficile (red lines), while cholate derivatives promote germination when combined with amino acid co-germinants like glycine (dotted line). Shen et al., 2015. correlation between CDI resistance, the presence of C. scindens and elevated levels of secondary bile acids have been observed [START_REF] Buffie | Precision microbiome reconstitution restores bile acid mediated resistance to Clostridium difficile[END_REF]. Combined with glycine, CAderived bile acids such as taurocholate promote germination of C. difficile spores (Sorg and Sonenshein 2008). By contrast, CDCA inhibits spore germination and growth of C. difficile vegetative cells (Sorg andSonenshein 2008, 2009), as well as the primary and secondary bile salt muricholates, which are present only in mice (Francis et al. 2013b). While deoxycholate found in humans and mice is a potent germinant, it is toxic, as the other secondary bile salts, for C. difficile vegetative cells (Sorg andSonenshein 2008, Francis et al. 2013b). Therefore, in a healthy mouse or human host, intestinal microbiota promotes the presence of a small amount of primary bile salts and a high amount of secondary bile salts in the colon (Theriot andYoung 2014, Buffie et al. 2015). This bile acids ratio leads a decrease in C. difficile spore germination and inhibition of vegetative cell growth (Figure 4) [START_REF] Theriot | Antibiotic-Induced Alterations of the Gut Microbiota Alter Secondary Bile Acid Production and Allow for Clostridium difficile Spore Germination and Outgrowth in the Large Intestine[END_REF].

Antibiotic usage is the main risk factor for CDI [START_REF] Mcfee | Clostridium difficile: emerging public health threat and other nosocomial or hospital acquired infections[END_REF][START_REF] Buffie | Profound alterations of intestinal microbiota following a single dose of clindamycin results in sustained susceptibility to Clostridium difficile-induced colitis[END_REF]) with the highest risk associated to the large spectrum antibiotics clindamycin, cephalosporins, penicillins, and fluoroquinolones [START_REF] Slimings | Antibiotics and hospital-acquired Clostridium difficile infection: update of systematic review and meta-analysis[END_REF]Riley 2014, Spigaglia 2016). Indeed, in response to antibiotic treatment, the composition of the intestinal microbiota is strongly and differently altered (Theriot andYoung 2014, Theriot et al. 2016). Treatment of mice with an antibiotic cocktail (ampicillin, gentamicin, metronidazole, neomycin and vancomycin) resulted in a decreased abundance of the Firmicutes phylum and increased persistence of Bacteroidetes and Proteobacteria [START_REF] Hill | Metagenomic analyses reveal antibiotic-induced temporal and spatial changes in intestinal microbiota with associated alterations in immune cell homeostasis[END_REF]. Other study, reported that after antibiotic treatment the gut community was dominated by the Proteobacteria phylum, Enterobacteriaceae family, whereas the Bacteroidetes and Firmicutes phyla only made up a small portion of the total population [START_REF] Antonopoulos | Reproducible community dynamics of the gastrointestinal microbiota following antibiotic perturbation[END_REF]. In humans, studies defining the impact of antibiotherapy in the structure of the gastrointestinal tract are more limited.

Antharam et al in 2013, compared fecal microbiota of healthy subjects and subjects with C. difficile-negative nosocomial diarrhea (CDN) or CDI. A decrease in bacterial diversity and species richness including Ruminococcaceae, Lachnospiracea and butyrogenic bacteria was observed in the fecal microbiota of CDN and CDI patients compared to healthy controls [START_REF] Antharam | Intestinal dysbiosis and depletion of butyrogenic bacteria in Clostridium difficile infection and nosocomial diarrhea[END_REF]. A more recent study comparing the differences in gut microbiome among CDI patients, asymptomatic carriers and healthy individuals showed an overabundance of Proteobacteria in the two first groups, and a decrease in the abundance of Bacteroidetes and Firmicutes, especially butyrate producers was observed [START_REF] Zhang | Insight into alteration of gut microbiota in Clostridium difficile infection and asymptomatic C. difficile colonization[END_REF].

Antibiotic treatment results also in changes in the pools of certain metabolites. Among bile acids, Giel et al demonstrated that primary bile acids are the ones that predominant in the

I.3. Epidemiology

C. difficile is a major cause of intestinal infection and diarrhea in individuals following antibiotic therapy (Rupnik et al. 2009[START_REF] Smits | Clostridium difficile infection[END_REF]. It is estimated that 7 to 17% of hospitalized patients carry toxinogenic strains of C. difficile [START_REF] Poutanen | Clostridium difficile-associated diarrhea in adults[END_REF]. CDI epidemiology has dramatically changed since the beginning of the 2000s. In the USA, between 2001 and 2010, the incidence of CDI among hospitalized adults approximately doubled [START_REF] Reveles | The rise in Clostridium difficile infection incidence among hospitalized adults in the United States: 2001-2010[END_REF]. C. difficile has become a major healthcare problem being responsible for half a million infections and 29.000 deaths each year, with a mortality rate estimated at 9.3% [START_REF] Lessa | Burden of Clostridium difficile infection in the United States[END_REF]. In Europe, the estimated number of cases is 124.000 per year [START_REF] Zarb | The European Centre for Disease Prevention and Control (ECDC) pilot point prevalence survey of healthcare-associated infections and antimicrobial use[END_REF]. However, CDI incidence is likely underestimated: on average, ~80 cases of CDI are missed per hospital per year in Europe [START_REF] Davies | Underdiagnosis of Clostridium difficile across Europe: the European, multicentre, prospective, biannual, point-prevalence study of Clostridium difficile infection in hospitalised patients with diarrhoea (EUCLID)[END_REF]. This is likely a result of a combination of factors such as the awareness of CDI among physicians and the type of methods/algorithm for CDI diagnosis implemented. In North America and in Europe, CDI outbreaks associated with increased severity of the disease and higher mortality rates are mainly linked to the emergence of strains belonging to the 027 ribotype and, to a lesser extent the 078 ribotype [START_REF] Cartman | The emergence of 'hypervirulence' in Clostridium difficile[END_REF][START_REF] Smits | Clostridium difficile infection[END_REF]). Potential mechanisms have been pointed for the increased severity of infections associated with 027 ribotype strains: increased toxin production [START_REF] Warny | Toxin production by an emerging strain of Clostridium difficile associated with outbreaks of severe disease in North America and Europe[END_REF], production of a binary toxin [START_REF] Cowardin | The binary toxin CDT enhances Clostridium difficile virulence by suppressing protective colonic eosinophilia[END_REF], fluoroquinolone resistance [START_REF] He | Emergence and global spread of epidemic healthcare-associated Clostridium difficile[END_REF], increased efficiency of sporulation [START_REF] Merrigan | Human hypervirulent Clostridium difficile strains exhibit increased sporulation as well as robust toxin production[END_REF]) and higher correlation with recurrent infection [START_REF] Marsh | Association of relapse of Clostridium difficile disease with BI/NAP1/027[END_REF]. The duration of hospital stay resultant of CDI range from 2.7 to 21.3 days for hospital-acquired CDI [START_REF] Gabriel | Hospitalization stay and costs attributable to Clostridium difficile infection: a critical review[END_REF].

Approximately 3%-4% of healthy adults are asymptomatically colonized by C. difficile. High carriage rates (25-35%) are reported in infants (< 1 year of age) and the rate drops to 15% for ages 1-8 years [START_REF] Enoch | Clostridium difficile in children: colonisation and disease[END_REF]. However, infants are frequently asymptomatic carriers of C. difficile. This is thought to be related to an immature microbiome and immune system [START_REF] Koenig | Succession of microbial consortia in the developing infant gut microbiome[END_REF][START_REF] Backhed | Dynamics and Stabilization of the Human Gut Microbiome during the First Year of Life[END_REF], protective maternal antibodies transferred in breast milk and/or the inability of the toxins to interact with the receptors and cause disease [START_REF] Eglow | Diminished Clostridium difficile toxin A sensitivity in newborn rabbit ileum is associated with decreased toxin A receptor[END_REF][START_REF] Rousseau | Clostridium difficile carriage in healthy infants in the community: a potential reservoir for pathogenic strains[END_REF]. Recent epidemiological studies have shown increased rates of community-associated C. difficile disease, affecting groups not previously at risk (such as younger patients and patients who had no exposure to antibiotics in the 12 weeks before infection) [START_REF] Khanna | The epidemiology of community-acquired Clostridium difficile infection: a population-based study[END_REF]). In addition, C. difficile is frequently encountered in a diverse range of animals that includes both pets and those destinated for meat production such as pork, veal and horse [START_REF] Gould | Clostridium difficile in food and domestic animals: a new foodborne pathogen?[END_REF][START_REF] Alvarez-Perez | Faecal shedding of antimicrobial-resistant Clostridium difficile strains by dogs[END_REF][START_REF] Warriner | Dissemination of Clostridium difficile in food and the environment: Significant sources of C. difficile community-acquired infection?[END_REF]. To date, direct transmission of C. difficile from animals, food or the environment to humans is not clear. However, a recent study reported that asymptomatic farmers and their pigs can be colonized with clonal isolates of C. difficile 078, showing evidences that spread between animals and humans might occur [START_REF] Knetsch | Whole genome sequencing reveals potential spread of Clostridium difficile between humans and farm animals in the Netherlands, 2002 to 2011[END_REF]. The increase incidence of community acquired C. difficile infection and the possible zoonotic transfer are a raising concern [START_REF] Smits | Clostridium difficile infection[END_REF]) and suggest additional and unrecognized risk factors for CDI.

I.4. Treatments of CDI

I.4.a. Conventional treatments

Initial therapy for CDI includes stopping the antibiotic therapy responsible for the emergence of C. difficile, if possible. If symptoms persist or worsen, antibiotic treatment against C. difficile is necessary. Metronidazole or vancomycin are used depending on the illness severity and co-morbidities. Metronidazole is the first line agent in the treatment of patients with mild-to-moderate CDI. Vancomycin is recommended for severe or complicated infections [START_REF] Surawicz | Guidelines for diagnosis, treatment, and prevention of Clostridium difficile infections[END_REF][START_REF] Debast | European Society of Clinical Microbiology and Infectious Diseases: update of the treatment guidance document for Clostridium difficile infection[END_REF]. Besides both antibiotics show similar cure rates in patients with mild-to-moderate infection, vancomycin is reserved for severe cases due to the growing concerns about antimicrobial-resistant microorganisms. In case of severe disease, vancomycin has higher cure rate [START_REF] Kazanowski | Clostridium difficile: epidemiology, diagnostic and therapeutic possibilities-a systematic review[END_REF]. One of the most challenging aspects of treating CDI is the recurrence of disease after a successful therapy. This makes recurrent CDI itself a significant risk factor. Recurrence can be caused by the initial infective strain or by spore reinfection from another strain acquired from the environment [START_REF] Barbut | Epidemiology of recurrences or reinfections of Clostridium difficile-associated diarrhea[END_REF]. The risk increases significantly with each episode of recurrence: 20-30% after a first episode and up to 60% after a first recurrence [START_REF] Shields | Recurrent Clostridium difficile infection: From colonization to cure[END_REF]. Treatment failures and recurrence of infection rates associated with the use of metronidazole and vancomycin were observed in the last years [START_REF] Vardakas | Treatment failure and recurrence of Clostridium difficile infection following treatment with vancomycin or metronidazole: a systematic review of the evidence[END_REF].

Therefore, other antibiotics such as rifampicin and fidaxomicin have been proposed to treat CDI. Fidaxomicin became available for the treatment of CDI in 2011. This antibiotic has equivalent cure rates to vancomycin but lower rate of CDI recurrence (13% vs 25%) with strains other than 027 rybotype [START_REF] Louie | Fidaxomicin versus vancomycin for Clostridium difficile infection[END_REF][START_REF] Cornely | Treatment of first recurrence of Clostridium difficile infection: fidaxomicin versus vancomycin[END_REF]. Contrary to vancomycin that has bacteriostatic activity against C. difficile, fidaxomicin is bactericidal [START_REF] Kazanowski | Clostridium difficile: epidemiology, diagnostic and therapeutic possibilities-a systematic review[END_REF]). Furthermore, it has a narrower spectrum compared to vancomycin and metronidazole, and thus has a lower impact on the composition of the gut microbiota, in particular on Bacteroides species [START_REF] Louie | Fidaxomicin preserves the intestinal microbiome during and after treatment of Clostridium difficile infection (CDI) and reduces both toxin reexpression and recurrence of CDI[END_REF]. However, the higher costs of fidaxomicin treatment compared to other therapeutic regimens may restrict its use. Alternative antimicrobials have been studied, such as the glycopeptide antibiotic teicoplanin. This compound has activity against Gram-positive anaerobes including C. difficile. A clinical trial has shown that teicoplanin was as effective as vancomycin and metronidazole for CDI.

However, the higher cost associated with its use made it as a drug used for the treatment of repeated recurrences of CDI [START_REF] Venugopal | Current state of Clostridium difficile treatment options[END_REF].

I.4.b. Faecal microbiota transplantation

The presence of C. difficile, either as an asymptomatic colonizer or as a pathogen, is associated with reduced microbiota diversity [START_REF] Zhang | Insight into alteration of gut microbiota in Clostridium difficile infection and asymptomatic C. difficile colonization[END_REF]. Similarly, patients with multiple episodes of CDI have a markedly reduced microbial diversity when compared to patients that suffered only a single episode of CDI [START_REF] Chang | Decreased diversity of the fecal Microbiome in recurrent Clostridium difficileassociated diarrhea[END_REF]. Thus, restoration of the complexity and diversity of the gut microbiota is required for C. difficile colonization resistance. Faecal microbiota transplantation (FMT) is a highly effective therapy that has gained much attention in the last years to treat recurrent CDI. The principle basis of FMT is that stools from a healthy donor contain a stable, viable, diverse and normal microbiota community. When introduced into the gut of a patient with CDI, they can restore their disrupted microbiota [START_REF] Brandt | Fecal transplantation for the treatment of Clostridium difficile infection[END_REF]. Studies have shown that FMT resolves recurrent CDI in more than 90% of the cases [START_REF] Van Nood | Duodenal infusion of donor feces for recurrent Clostridium difficile[END_REF], Kelly et al. 2015b). A recent study on 18 patients shown a CDI healing rate of 94% after FMT [START_REF] Konturek | Successful therapy of Clostridium difficile infection with fecal microbiota transplantation[END_REF]. Following successful transplantation, Bacteroidetes increased and Proteobacteria decreased in fecal microbiota [START_REF] Seekatz | Recovery of the gut microbiome following fecal microbiota transplantation[END_REF]. Interestingly, a recent study of five CDI patients showed that transfer of sterile filtrates from donor stool rather than fecal microbiota, was sufficient to eliminate symptoms. In most treated patients, the faecal filtrated transplantation was associated with a reduction of enterobacteracea. These results suggest that bacterial components, such as metabolites or bacteriophages might be sufficient but they cannot define the exact mechanism of action [START_REF] Ott | Efficacy of Sterile Fecal Filtrate Transfer for Treating Patients With Clostridium difficile Infection[END_REF].

I.4.c. Immunotherapy

Since C. difficile TcdA and TcdB are the main factors responsible for CDI symptoms, most of the immune therapies that have been undertaken target these proteins. A monoclonal antibody raised against the toxin B, Zinplava, was recently approved by the FDA.

Introduction 20

Administration of Zinplava in combination with the standard antibacterial drugs (metronidazole, vancomycin or fidaxomicin) resulted in a lower rate of CDI recurrence [START_REF] Wilcox | Bezlotoxumab for Prevention of Recurrent Clostridium difficile Infection[END_REF]). An antibody raised against toxin A (Actoxumab) was also developed but it provided no additional benefit over Zinplava. Active immunization has been also tested. An oral vaccination trial has been performed with spores of Bacillus subtilis expressing TcdA peptide repeats, which confer protection in a hamster model after challenge with the C. difficile 630 strain (Permpoonpattana et al. 2011a). C. difficile colonization can lead to asymptomatic carriage, or a wide range of symptoms, from mild diarrhea, abdominal pain, fever and leucocytosis, to more severe symptoms such as pseudomembranous colitis, toxic megacolon, bowel perforation, sepsis and death [START_REF] Dawson | Clostridium difficile--a continually evolving and problematic pathogen[END_REF], Rupnik et al. 2009). These symptoms are mainly caused by the large toxins TcdA and TcdB [START_REF] Voth | Clostridium difficile toxins: mechanism of action and role in disease[END_REF][START_REF] Aktories | Clostridium difficile Toxin Biology[END_REF][START_REF] Chandrasekaran | The role of toxins in Clostridium difficile infection[END_REF]. The expression of the genes encoding TcdA (308 kDa) and TcdB (270 kDa) toxins is induced during stationary growth phase when nutrients become limited [START_REF] Hundsberger | Transcription analysis of the genes tcdA-E of the pathogenicity locus of Clostridium difficile[END_REF]. TcdA and TcdB have four domains: an N-terminal glucosyltransferase (GDT) domain that inactivates members of the Rho family of small GTPases (small regulatory proteins of the eukaryote actin cell cytoskeleton) by glucosylation; an autoprocessing (ADP) domain; a delivery/pore forming domain and a receptor-binding (CROPS) domain (Figure 5A). These domains contribute to a multistep mechanism of intoxication (Figure 5B). Once released, TcdA and TcdB bind to specific receptors on intestinal epithelial cells and are internalized by specific endocytic pathways. TcdA surface receptors are not clearly identified, however, TcdA is able to bind to glycoproteins such as Gp96 in the human colonocytes [START_REF] Na | gp96 is a human colonocyte plasma membrane binding protein for Clostridium difficile toxin A[END_REF]) and to the sucrose-isomaltase located in the brush border of small intestines [START_REF] Pothoulakis | Rabbit sucrase-isomaltase contains a functional intestinal receptor for Clostridium difficile toxin A[END_REF]. After the identification of CSPG4 as a receptor for TcdB, two other reports were published showing that NECTIN3 (also termed poliovirus receptor-like protein (PVRL3)) and frizzled proteins 1, 2 and 7, function as colonic epithelial receptors for TcdB [START_REF] Lafrance | Identification of an epithelial cell receptor responsible for Clostridium difficile TcdB-induced cytotoxicity[END_REF][START_REF] Tao | Frizzled proteins are colonic epithelial receptors for C. difficile toxin B[END_REF]. The low pH in the endosome triggers a conformational change in the toxin delivery domain resulting in pore Both tcdA and tcdB genes are encoded on the 19.6-kb pathogenicity locus (PaLoc), together with tcdR, tcdC and tcdE genes. TcdR is an alternative sigma factor required for the transcription of tcdA, tcdB and tcdR itself. TcdC is an anti-sigma factor which sequesters TcdR preventing the transcription of the toxin genes. TcdE is a "holin" type bacteriophage protein required for toxin secretion.

II. Virulence factors and colonization in

tcdE tcdB tcdR tcdC tcdA

formation that allows translocation of the GDT across the endosome membrane. GTD is autoprocessed and released to the cytosol where it catalyzes the glucosylation and hence the inactivation of Rho-and Ras-GTPases rendering them inactive (Just et al. 1995a, Just et al. 1995b). Consequences of this are disorganization of the cytoskeleton, cell rounding and eventually cell death by apoptosis (Figure 5B) [START_REF] Warny | p38 MAP kinase activation by Clostridium difficile toxin A mediates monocyte necrosis, IL-8 production, and enteritis[END_REF][START_REF] Nottrott | Clostridium difficile toxin Ainduced apoptosis is p53-independent but depends on glucosylation of Rho GTPases[END_REF]. TcdB also induces necrosis by triggering the production of reactive oxygen species (ROS) through the assembly of the NADPH oxidase (NOX) complex on endosomes [START_REF] Farrow | Clostridium difficile toxin B-induced necrosis is mediated by the host epithelial cell NADPH oxidase complex[END_REF].

High levels of ROS promote cellular necrosis likely through DNA damage and protein oxidation. Additionally C. difficile toxins stimulate the production of pro-inflammatory cytokines by colonic epithelial cells which lead to an innate inflammatory response with neutrophil recruitment [START_REF] Voth | Clostridium difficile toxins: mechanism of action and role in disease[END_REF]. Inflammation of the colon occurs with massive fluid loss into the large intestine, clinically presenting as diarrhea [START_REF] Voth | Clostridium difficile toxins: mechanism of action and role in disease[END_REF][START_REF] Jank | Rho-glucosylating Clostridium difficile toxins A and B: new insights into structure and function[END_REF][START_REF] Chandrasekaran | The role of toxins in Clostridium difficile infection[END_REF]. Despite the development of genetic tools, the relative contribution of the TcdA and TcdB in the pathogenesis of C. difficile is still relatively controversial. The combined deletion of both TcdA and TcdB completely abolish in vivo virulence [START_REF] Kuehne | The role of toxin A and toxin B in Clostridium difficile infection[END_REF][START_REF] Kuehne | Importance of toxin A, toxin B, and CDT in virulence of an epidemic Clostridium difficile strain[END_REF]. In order to undersant the respective role of each toxin in the virulence process, a study performed in 2009 showed that TcdB, unlike TcdA, is essential for the virulence of C. difficile [START_REF] Lyras | Toxin B is essential for virulence of Clostridium difficile[END_REF]. However, another study demonstrated that the presence of either TcdA or TcdB alone is sufficient to cause death in the hamster [START_REF] Kuehne | The role of toxin A and toxin B in Clostridium difficile infection[END_REF][START_REF] Kuehne | Importance of toxin A, toxin B, and CDT in virulence of an epidemic Clostridium difficile strain[END_REF], suggesting that both toxins are capable of inducing disease. In agreement, some strains isolated from patients with classical symptoms of CDIs have been found to produce either TcdA or TcdB [START_REF] Carter | The role of toxin A and toxin B in the virulence of Clostridium difficile[END_REF][START_REF] Monot | Clostridium difficile: New Insights into the Evolution of the Pathogenicity Locus[END_REF]. Nevertheless, it appears that TcdB toxin is more important in the activation of the host immune system and the inflammatory response [START_REF] Carter | Defining the Roles of TcdA and TcdB in Localized Gastrointestinal Disease, Systemic Organ Damage, and the Host Response during Clostridium difficile Infections[END_REF].

II.1.b. Regulation of tcdA and tcdB expression

The genes tcdA and tcdB encoding respectively TcdA and TcdB, are located in the 19.6-kb pathogenicity locus (PaLoc) (Figure 6) [START_REF] Braun | Definition of the single integration site of the pathogenicity locus in Clostridium difficile[END_REF]. In most non-toxigenic strains, the PaLoc is replaced by a 75-115 nucleotide non-coding sequence, while a 7.2-kb sequence of unknown function has been also found in some Tox -strains [START_REF] Braun | Definition of the single integration site of the pathogenicity locus in Clostridium difficile[END_REF][START_REF] Monot | Clostridium difficile: New Insights into the Evolution of the Pathogenicity Locus[END_REF]. The PaLoc also carries three aditional genes: tcdE, tcdR and tcdC (Figure 6). TcdE is a holin-like protein whose pore-forming activity is involved in the release of toxin [START_REF] Govind | Secretion of Clostridium difficile toxins A and B requires the holinlike protein TcdE[END_REF]. However the role of TcdE in the toxin secretion has been controversial [START_REF] Govind | Secretion of Clostridium difficile toxins A and B requires the holinlike protein TcdE[END_REF][START_REF] Olling | Release of TcdA and TcdB from Clostridium difficile cdi 630 is not affected by functional inactivation of the tcdE gene[END_REF], Govind et al. 2015, Monot et . Indeed, Olling et al., showed that toxin secretion was not affected by inactivation of the tcdE gene [START_REF] Olling | Release of TcdA and TcdB from Clostridium difficile cdi 630 is not affected by functional inactivation of the tcdE gene[END_REF]). Recently, it has been demonstrated that in response to specific environmental conditions toxin release is also mediated by an autolysin (Cwp19) and is associated by autolysis [START_REF] Wydau-Dematteis | Cwp19 Is a Novel Lytic Transglycosylase Involved in Stationary-Phase Autolysis Resulting in Toxin Release in Clostridium difficile[END_REF]). In addition they showed that both TcdE and bacteriolysis are coexisting mechanism for toxin release according to the growth condition. TcdR-encoding gene is positively regulated by σ D , the σ factor controlling flagellum synthesis [START_REF] El Meouche | Characterization of the SigD regulon of C. difficile and its positive control of toxin production through the regulation of tcdR[END_REF][START_REF] Mckee | The second messenger cyclic Di-GMP regulates Clostridium difficile toxin production by controlling expression of sigD[END_REF]. TcdR is an alternative sigma factor that interacts with the core enzyme of the RNA polymerase (RNAP) to specifically recognize and activate transcription of toxin genes. TcdR also positively controls its own expression [START_REF] Mani | Regulation of toxin synthesis in Clostridium difficile by an alternative RNA polymerase sigma factor[END_REF]. Finally, TcdC is an anti-sigma factor that negatively regulates toxin gene expression by interfering with the ability of the TcdRcontaining RNAP to recognize the tcdA and tcdB promoters [START_REF] Matamouros | Clostridium difficile toxin expression is inhibited by the novel regulator TcdC[END_REF][START_REF] Dupuy | Clostridium difficile toxin synthesis is negatively regulated by TcdC[END_REF][START_REF] Carter | The anti-sigma factor TcdC modulates hypervirulence in an epidemic BI/NAP1/027 clinical isolate of Clostridium difficile[END_REF]. Toxin expression is regulated by growth conditions and metabolic changes (Figure 7) [START_REF] Martin-Verstraete | The Regulatory Networks That Control Clostridium difficile Toxin Synthesis[END_REF]. In fact, the presence of rapidly metabolizable carbon sources imported by the phosphotransferase-like system (PTS) such as glucose, fructose and mannitol represses the expression of toxin encoding genes [START_REF] Dupuy | Regulated transcription of Clostridium difficile toxin genes[END_REF]. These repression is mediated by CcpA, the global regulator of the carbon catabolic repression system [START_REF] Deutscher | How phosphotransferase system-related protein phosphorylation regulates carbohydrate metabolism in bacteria[END_REF]. CcpA interacts directly with the promoter region of several PaLoc genes, with the strongest afinity for the promoter region of tcdR [START_REF] Antunes | Global transcriptional control by glucose and carbon regulator CcpA in Clostridium difficile[END_REF]. The global transcriptional repressor CodY, which is involved in the adaptive response to nutrient deficiencies during entry in stationary phase, is responsible for the repression of toxin genes during the exponential growth phase [START_REF] Dineen | Repression of Clostridium difficile toxin gene expression by CodY[END_REF]). The affinity of CodY to its targets genes is increased in the presence of branched-chain aminoacids (BCAA) and GTP (Ratnayake-Lecamwasam et al. 2001, Shivers and[START_REF] Shivers | Activation of the Bacillus subtilis global regulator CodY by direct interaction with branched-chain amino acids[END_REF].

In these conditions, CodY negatively regulates toxin gene expression by binding to the tcdR promoter region [START_REF] Dineen | Repression of Clostridium difficile toxin gene expression by CodY[END_REF]. Finally, in addition to the BCAAs, amino acids like proline, glycine and cysteine have a significant inhibitory effect on toxin production [START_REF] Karlsson | Suppression of toxin production in Clostridium difficile VPI 10463 by amino acids[END_REF][START_REF] Dubois | Control of Clostridium difficile Physiopathology in Response to Cysteine Availability[END_REF]. The impact of cysteine on tcdA and tcdB expression is mediated by the σ factor σ L . σ L is involved in the metabolism and virulence of Grampositive bacteria [START_REF] Okada | The sigma factor RpoN (sigma54) is involved in osmotolerance in Listeria monocytogenes[END_REF]). However, the absence of a consensus sequence for σ L upstream of the PaLoc genes suggests an indirect effect on toxin expression. In C. difficile, σ L controls the degradation of cysteine into pyruvate and sulfides, compounds that negatively affect the expression of toxins when they are added to the culture medium [START_REF] Dubois | Control of Clostridium difficile Physiopathology in Response to Cysteine Availability[END_REF]. Since toxin production and sporulation take place during transition growth phase, the The monomeric form of CDTb binds to its cell surface receptor, LSR. Thereafter, CDTb undergoes proteolytic activation and oligomerization on the cell surface. This process might also occur before binding. The oligomeric prepore binds CDTa, and the receptor-CDT complex is subsequently internalized into cells. As the endosome matures, the acidic pH triggers conformational changes in CDTb, resulting in membrane insertion and formation of a pore through which CDTa is translocated into the cytosol. Translocation of CDTa is facilitated by heat shock protein 70 (Hsp70), Hsp90 and cyclophilin A (CypA). In the cytosol, CDTa ADP-ribosylates actin at arginine-177. ADP-ribosylation of actin results in the eventual breakdown of the actin cytoskeleton and cytopathic cell rounding, and the formation of microtubule-based protrusions that enhance bacterial adherence. [START_REF] Chandrasekaran | The role of toxins in Clostridium difficile infection[END_REF] connection between the two mechanisms was studied [START_REF] Underwood | Characterization of the sporulation initiation pathway of Clostridium difficile and its role in toxin production[END_REF]). The inactivation of sigH, coding for the key σ factor of the phase transition, results in increased expression of toxin encoding genes (Saujet et al. 2011). In addition, inactivation of the master regulator of sporulation spo0A, results in the upregulation of tcdA gene [START_REF] Underwood | Characterization of the sporulation initiation pathway of Clostridium difficile and its role in toxin production[END_REF][START_REF] Pettit | Functional genomics reveals that Clostridium difficile Spo0A coordinates sporulation, virulence and metabolism[END_REF]. The absence of the binding domain of Spo0A in the promoter region of toxin genes, suggests that Spo0A controls indirectly toxin genes transcription probably via one of the other factors that control toxin gene expression [START_REF] Dineen | Integration of metabolism and virulence by Clostridium difficile CodY[END_REF], Antunes et al. 2011[START_REF] Bouillaut | Proline-dependent regulation of Clostridium difficile Stickland metabolism[END_REF]). More recently, new links between sporulation and toxin synthesis have been demonstrated (Figure 7). RstA, a Rap-like protein, positively controls sporulation and negatively regulates toxin production (Edwards et al. 2016b). RstA negatively influences toxin production by repressing the transcription of sigD. Indeed, the inactivation of rstA leads to an increased expression of sigD and of σ D -controlled genes (Edwards et al. 2016b).

II.2. The binary toxin CDT

Some C. difficile strains, as those of PCR ribotypes 027 and 078, also produce a third toxin called C. difficile transferase CDT [START_REF] Aguilar | KinD is a checkpoint protein linking spore formation to extracellular-matrix production in Bacillus subtilis biofilms[END_REF]. CDT belongs to the family of binary actin ADP-ribosylating toxins and is encoded by two genes, cdtA and cdtB located on a 6.2-kb chromosomal region (named the Cdt locus or CdtLoc). A third gene, cdtR, located in the same locus, encodes a regulator that positively controls CDT production [START_REF] Carter | Binary toxin production in Clostridium difficile is regulated by CdtR, a LytTR family response regulator[END_REF]) and in some C. difficile clinical strains regulates TcdA and TcdB production [START_REF] Lyon | CdtR Regulates TcdA and TcdB Production in Clostridium difficile[END_REF]).

CDT-negative toxigenic strains typically contain a 2-kb deletion within the CdtLoc [START_REF] Stare | Variant forms of the binary toxin CDT locus and tcdC gene in Clostridium difficile strains[END_REF]). As binary toxin, the CDTb component (100 kDa) binds to its membrane receptor, the lipolysis-stimulated lipoprotein receptor (LSR) (Figure 8). Then, proteolytic activation of CDTb occurs, which is necessary for oligomerization and formation of a heptamer.

Alternatively, the monomeric form of CDTb may undergo proteolytic activation and oligomer formation before binding to LSR. Then, the catalytic component CDTa (50 kDa) interacts with CdtB and the LSR-CDT complex is endocytosed reaching an acidic endossomal compartment. The acidic pH of the endosome triggers conformational changes in CDTb, resulting in membrane insertion and formation of a pore that allows CDTa translocation to the cytosol. This process is supported by cytosolic chaperones such as the heat shock protein 70 (Hsp70), Hsp90 and cyclophilin A (CypA). In the cytosol, CTDa ADP ribosylates monomeric actin and this inhibits actin polymerization. Furthermore, depolymerization of actin favors the formation of microtubule-based protrusions that enhance adherence and colonization of C.

difficile (Figure 8) [START_REF] Schwan | Clostridium difficile toxin CDT induces formation of microtubule-based protrusions and increases adherence of bacteria[END_REF][START_REF] Chandrasekaran | The role of toxins in Clostridium difficile infection[END_REF][START_REF] Aguilar | KinD is a checkpoint protein linking spore formation to extracellular-matrix production in Bacillus subtilis biofilms[END_REF].

The environmental signals that regulate CDT production and the mechanism of toxin secretion have not yet been studied.

II.3. Other virulence factors: the colonization factors

Like many enteric pathogens, the vegetative cells of C. difficile are likely associate with the intestinal mucosa to begin the process of host colonization and establish infection.

Therefore, adherence and motility must be important factors contributing to colonization efficiency and virulence of this pathogen [START_REF] Awad | Clostridium difficile virulence factors: Insights into an anaerobic spore-forming pathogen[END_REF]. Studies have shown that C.

difficile vegetative cells are able to bind to intestinal epithelial cells and, in vitro, interact with apical microvilli of differentiated Caco-2 cells [START_REF] Eveillard | Identification and characterization of adhesive factors of Clostridium difficile involved in adhesion to human colonic enterocyte-like Caco-2 and mucus-secreting HT29 cells in culture[END_REF][START_REF] Karjalainen | Cloning of a genetic determinant from Clostridium difficile involved in adherence to tissue culture cells and mucus[END_REF][START_REF] Cerquetti | Binding of Clostridium difficile to Caco-2 epithelial cell line and to extracellular matrix proteins[END_REF]. Moreover, the vegetative cells can also adhere in vivo to the mucosal epithelium of hamsters or mice [START_REF] Borriello | Mucosal association by Clostridium difficile in the hamster gastrointestinal tract[END_REF]. The cell surface of C. difficile is covered by adhesion factors such as the S-layer proteins that facilitate binding to epithelial cells and to components of the extracellular matrix such as fibrinogen, laminin, fibronectin and collagen [START_REF] Cerquetti | Binding of Clostridium difficile to Caco-2 epithelial cell line and to extracellular matrix proteins[END_REF][START_REF] Spigaglia | Surface-layer (S-layer) of human and animal Clostridium difficile strains and their behaviour in adherence to epithelial cells and intestinal colonization[END_REF]. It was shown that adherence of C.

difficile to epithelial cells is greatly reduced on cell monolayers when they are pre-incubated with purified SlpA proteins, the major components of the S-layer [START_REF] Merrigan | Surface-layer protein A (SlpA) is a major contributor to host-cell adherence of Clostridium difficile[END_REF].

Several other proteins located at the cell surface play a role in adherence to intestinal epithelial cells: i.e., cell wall protein, such as Cwp66 with adhesive properties [START_REF] Waligora | Characterization of a cell surface protein of Clostridium difficile with adhesive properties[END_REF]; the fibronectin-binding protein Fbp68 [START_REF] Hennequin | Identification and characterization of a fibronectin-binding protein from Clostridium difficile[END_REF]; the lipoprotein CD0873 involved in adhesion to Caco2 cells [START_REF] Kovacs-Simon | Lipoprotein CD0873 is a novel adhesin of Clostridium difficile[END_REF]) and the flagella composed of the flagellin FliC and the flagellar cap protein FliD, which are involved in mucus attachment (Tasteyre et al. 2001), and (iv). The role of FliC and FliD in adhesion to Caco-2 cells differs according to C. difficile strains. Indeed, the inactivation of fliC and fliD genes in the 630Δerm strain increased cell adhesion and virulence [START_REF] Dingle | Mutagenic analysis of the Clostridium difficile flagellar proteins, FliC and FliD, and their contribution to virulence in hamsters[END_REF][START_REF] Baban | The role of flagella in Clostridium difficile pathogenesis: comparison between a non-epidemic and an epidemic strain[END_REF]) while the same inactivation in strain R20291 (ribotype 027) decreases adhesion [START_REF] Baban | The role of flagella in Clostridium difficile pathogenesis: comparison between a non-epidemic and an epidemic strain[END_REF]. In unstressed cells and during the exponential growth phase, the anti-sigma factor, RsbW, phosporylates the anti-anti sigma, RsbV, and sequesters σ B rendering it inactive. During stationary growth phase of after stress exposure, the non-phosphorylated form of RsbV interacts with RsbW, allowing σ B release, its association with the RNA polymerase and transcription of its targets genes.

I. Stresses encountered during gut infection

Pathogenic bacteria experience diverse stresses while spreading to and colonizing new hosts to cause infection. Enteric pathogens when ingested, must survive to the acidic pH of the stomach [START_REF] Foster | When protons attack: microbial strategies of acid adaptation[END_REF]. Within the intestinal lumen, bacteria encounters other stresses such as bile salts, free fatty acids, pH variations, hyperosmolarity, antibiotics, antimicrobial peptides and changing oxygen (O 2 ) tensions [START_REF] Guiney | Regulation of bacterial virulence gene expression by the host environment[END_REF]. Moreover, host inflammatory responses recruit phagocytic cells which subjet pathogens to oxidative and nitrosative stress [START_REF] Fang | Antimicrobial reactive oxygen and nitrogen species: concepts and controversies[END_REF]). To survive to this hostile environment, bacteria have to develop mechanisms to sense these stresses and trigger appropriate responses that can allow their survival. Bacteria respond by altering their transcriptome and proteome in an adaptative manner.

II. σ B , the alternative sigma factor of general stress response

The most effective response to a wide range of stresses and starvation conditions in several Firmicutes is mediated by σ B , the alternative sigma factor of general stress response (Hecker et al. 2007). The σ B -dependent responses are conserved in the genera Bacillus, Staphylococcus, Listeria and in a few clostridia. The sigB gene is usually associated with genes encoding the anti-anti-sigma factor, RsbV, and the sigma-factor, RsbW involved in the control of σ B activity through a well-conserved post-translational mechanism, called partner switching (Alper et al. 1996).

II.1 σ B in B. subtilis

II.1.a The role of σ B and the mechanism of its activation

In B. subtilis, σ B , RsbV, and RsbW are encoded within an eight-gene operon that is expressed constitutively from a σ A -dependent promoter present upstream of rsbRA (Figure 9A) (Kalman et al. 1990, Wise andPrice 1995). However, an internal σ B -dependent promoter enhances expression of the four last genes of the operon (rsbV, rsbW, sigB, and rsbX) when σ B is active (Kalman et al. 1990, Benson andHaldenwang 1993b). σ B activity is controlled by a post-translational mechanism, in which interaction and activation of proteins are driven by

Low ATP

reversible threonine and serine phosphorylation (Figure 9B) (Yang et al. 1996, Hecker et al. 2007). In fact, the phosphorylated state of the anti-anti-sigma factor, RsbV, is the key element of σ B activation or silencing (Alper et al. 1996). In unstressed cells and during the exponential growth phase, the kinase RsbW, phosphorylates RsbV and sequesters as an anti-sigma factor, σ B thereby preventing its association with the RNA polymerase (Benson and Haldenwang 1993a). When cells are exposed to a stress or in the stationary phase, a signal is transmitted to specific PP2C phosphatase(s) that dephosphorylate(s) RsbV and allows interaction of RsbV with RsbW, leading to the release and activation of σ B (Figure 9B). The σ B -containing holozyme will ensure the recognition of appropriate promoters upstream of the controlled genes and their transcription.

II.1.b. The signal transduction pathway responsible for the activation of σ B

Stress signals triggering phosphatase activation and σ B release depend on the bacterial species. Stress stimuli include both high and low temperatures, salt, ethanol, acidic pH, hydrogen peroxide (H 2 O 2 ), nitric oxide (NO), irradiation with blue light, presence of antibiotics inducing cell wall stress, and starvation for glucose, phosphate or O 2 . The carbonyl cyanide m-chlorophenylhydrazone (CCCP), which cause a deprivation of ATP and GTP levels can also triggers σ B activation (Benson and Haldenwang 1993c, Boylan et al. 1993, Price et al. 2001, Helmann et al. 2003[START_REF] Mascher | Cell wall stress responses in Bacillus subtilis: the regulatory network of the bacitracin stimulon[END_REF][START_REF] Moore | Response of Bacillus subtilis to nitric oxide and the nitrosating agent sodium nitroprusside[END_REF], Zhang and Haldenwang 2005[START_REF] Avila-Perez | Blue light activates the sigmaB-dependent stress response of Bacillus subtilis via YtvA[END_REF][START_REF] Gaidenko | The blue-light receptor YtvA acts in the environmental stress signaling pathway of Bacillus subtilis[END_REF], Hecker et al. 2007[START_REF] Avila-Perez | Red light activates the sigmaB-mediated general stress response of Bacillus subtilis via the energy branch of the upstream signaling cascade[END_REF]. In B. subtilis, σ B can be activated via two main signaling pathways that respond to energy depletion or environmental stresses (Figure 10). For example, deficiency in glucose, phosphate, O 2 limitation or entry in stationary phase induces gene expression in a σ Bdependent manner. All these conditions lead to a decrease in the amount of intracellular ATP [START_REF] Maul | sigma B-dependent regulation of gsiB in response to multiple stimuli in Bacillus subtilis[END_REF][START_REF] Antelmann | Phosphate starvation-inducible proteins of Bacillus subtilis: proteomics and transcriptional analysis[END_REF], Hecker et al. 2007). Therefore, addition of CCCP or a high concentration of Mn 2+ or Mg 2+ (competitor ions used as a cofactor in ATP-generating reactions) that decreases the intracellular concentration of ATP increases the expression of σ B -controlled genes (Voelker et al. 1996a, Zhang andHaldenwang 2005). RsbP is an RsbVspecific phosphatase that together with RsbQ senses energy limitation (Figure 10A). In addition to its catalytic PP2C-phosphatase domain, RsbP contains a PAS domain essential for its activity (Vijay et al. 2000). PAS domains are known to control interaction with other proteins, to detect changes in the energy level of the cells, or to sense the redox potential, light or O 2 concentration. Two models have been proposed for the RsbP activation (Nadezhdin et al. 2011). In the first model, RsbQ converts a small molecule into a product delivered to the PAS domain of RsbP resulting in its conformational change. In the second model, RsbQ is directly associated with the PAS domain of RsbP and the transformation of a small molecule generates a conformational change in the PAS domain. In both cases, the conformational modification of the PAS domain activates the RsbP PP2C-phosphatase, which dephosphorylate RsbV~P, resulting in the activation of σ B [START_REF] Brody | Bypass suppression analysis maps the signalling pathway within a multidomain protein: the RsbP energy stress phosphatase 2C from Bacillus subtilis[END_REF], Nadezhdin et al. 2011). In B. subtilis, environmental stresses that activate σ B are recognized by a multicomponent protein complex, the stressosome, which transmit the signal to another PP2C-phosphatase, RsbU (Figure 10) (Voelker et al. 1996b, Yang et al. 1996). In growing cells, RsbU is inactive. Activation of RsbU is depending on the binding and stimulation by the kinase, RsbT. However, under growing conditions, RsbT is captured by its antagonist RsbS. Together with RsbS and RsbR, RsbT is part of the stressosome. Upon environmental stress, the kinase activity of RsbT is stimulated by an unknown mechanism, and the stressossome components, RsbR and RsbS, are phosphorylated. Their phosphorylation promotes the dissociation of the RsbS-RsbT complex, releasing RsbT from the stressosome (Figure 10B) (Yang et al. 1996) and allows its association with RsbU. When bound to RsbT, the RsbU phosphatase domain is activated leading to the dephosphorylation of RsbV~P and the activation of σ B . Finally, a third pathway of σ B activation occurs during continuous growth at low temperature [START_REF] Brigulla | Chill induction of the SigB-dependent general stress response in Bacillus subtilis and its contribution to lowtemperature adaptation[END_REF]. However this activation, which is RsbVindependent, is not yet well characterized.

II.2. Diversity of the mechanism of σ B activation in other Firmicutes

Interestingly, while the role of RsbV and RsbW in σ B activation is highly conserved, both signal and factors regulating σ B activation upstream of RsbV differ considerably among Firmicutes. RsbQ and RsbP involved in sensing-energy branch are confined to B. subtilis. In L. monocytogenes, the pathway of σ B activation by environmental stress such acidic pH, osmotic and heat stresses, is similar to that observed in B. subtilis with the presence of the RsbRST proteins and a stressosome (Becker et al. 1998[START_REF] Ferreira | Role of sigma(B) in heat, ethanol, acid, and oxidative stress resistance and during carbon starvation in Listeria monocytogenes[END_REF][START_REF] Smith | How does Listeria monocytogenes combat acid conditions?[END_REF]. Even if L. monocytogenes lacks RsbP and RsbQ, ATP depletion induces σ B activity in an RsbT-dependent manner indicating that environmental and energy signals seems to be detected by the same pathway in this bacterium (Chaturongakul andBoor 2004, 2006). In L. monocytogenes, as in B. subtilis (Figure 9A), the rsbVW-sigB-rsbX genes are transcribed by a σ B and rsbRSTU genes located upstream of rsbV are expressed from a σ A -dependent promoter (Wise andPrice 1995, Ferreira et al. 2004). In S. aureus, the stressosome components RsbR, RrbS and RsbT as well as the proteins involved in the energy pathway, RsbP and RsbQ, are missing. Accordingly, depletion of ATP or an ethanol stress, triggering σ B activation in B. subtilis, do not activate σ B in S. aureus (Pane-Farre et al. 2006). In S. aureus, the activation of σ B , which only involves RsbU, RsbV and RsbW (Giachino et al. 2001, Senn et al. 2005), is mainly activated upon entry into stationary phase or in the presence of alkaline stress (Giachino et al. 2001, Pane-Farre et al. 2006, Pane-Farre et al. 2009). Nevertheless, the mechanisms allowing the activation of RsbU in these conditions remain unknown. In this bacterium, the rsbUWV-sigB operon is transcribed from a σ A -dependent promoter located upstream of rsbU and a σ B -dependent promoter is present upstream of rsbV (Senn et al. 2005). In B. cereus group (B. cereus, B. anthracis, and B. thuringiensis), the absence of correlation between intracellular level of ATP and σ B activation suggest that σ B is not activated by energy depletion. In B. cereus, σ B is activated during heat or osmotic shock and after ethanol exposure (van Schaik et al. 2004). This bacterium uses another phosphatase protein, RsbY and a sensor protein RsbK different from RsbQ and RsbT suggesting a specific mechanism of sensing environmental stresses (van Schaik et al. 2004, van Schaik et al. 2005).

Despite differences in the sigB operon structure, all of these Firmicutes have a σ B -dependent positive-feedback loop that regulates the transcription of the sigB and rsb genes.

II.3. The σ B regulon

Large-scale transcriptomic and proteomic studies have been made to determine the genes controlled by σ B . The σ B regulon of B. subtilis, L. monocytogenes and S. aureus consists in approximately 150 genes positively regulated by σ B [START_REF] Petersohn | Global analysis of the general stress response of Bacillus subtilis[END_REF], Price et al. 2001[START_REF] Bischoff | Microarray-based analysis of the Staphylococcus aureus sigmaB regulon[END_REF], Pane-Farre et al. 2006[START_REF] Van Schaik | Identification of the sigmaB regulon of Bacillus cereus and conservation of sigmaB-regulated genes in low-GC-content gram-positive bacteria[END_REF][START_REF] Raengpradub | Comparative analysis of the sigma Bdependent stress responses in Listeria monocytogenes and Listeria innocua strains exposed to selected stress conditions[END_REF][START_REF] Oliver | Listeria monocytogenes {sigma}B has a small core regulon and a conserved role in virulence but makes differential contributions to stress tolerance across a diverse collection of strains[END_REF][START_REF] Nannapaneni | Defining the structure of the general stress regulon of Bacillus subtilis using targeted microarray analysis and random forest classification[END_REF][START_REF] Mujahid | Refinement of the Listeria monocytogenes sigmaB regulon through quantitative proteomic analysis[END_REF], Guldimann et al. 2016[START_REF] Mader | Staphylococcus aureus Transcriptome Architecture: From Laboratory to Infection-Mimicking Conditions[END_REF]. Some σ B -controlled genes are directly involved in the management of stresses, whereas others participate in various cellular processes such as metabolism, cell envelope homeostasis, biofilm formation and virulence (Guldimann et al. 2016). In addition, σ B also positively and negatively controls the expression of genes encoding many regulators (about 20 in B. subtilis) and then indirectly regulates expression of their target genes and the adaptive responses associated. For example, σ B transcribes the gene encoding the global regulator MgsR, which controls the expression of a sub-regulon involved in the oxidative stress response in B. subtilis [START_REF] Reder | The Spx paralogue MgsR (YqgZ) controls a subregulon within the general stress response of Bacillus subtilis[END_REF]). Among the MgsR targets is the transcriptional regulator NsrR involved in NO response resulting in a complex regulatory cascade.

In bacilli, σ B controls key functions involved in the response to heat, acid, starvation, NO or osmotic stress (Hecker et al. 2007[START_REF] Nannapaneni | Defining the structure of the general stress regulon of Bacillus subtilis using targeted microarray analysis and random forest classification[END_REF], Reder et al. 2012b). In Listeria, σ B participates in the responses to acid, oxidative, and energy stresses (Wiedmann et al. 1998[START_REF] Ferreira | Role of sigma(B) in heat, ethanol, acid, and oxidative stress resistance and during carbon starvation in Listeria monocytogenes[END_REF][START_REF] Chaturongakul | RsbT and RsbV contribute to sigmaB-dependent survival under environmental, energy, and intracellular stress conditions in Listeria monocytogenes[END_REF]. In S. aureus, σ B appears also to control cell wall metabolism, membrane transport and resistance to several clinically important antibiotics, such as methicillin and vancomycin (Pane-Farre et al. 2006[START_REF] Schulthess | Functional characterization of the sigmaB-dependent yabJ-spoVG operon in Staphylococcus aureus: role in methicillin and glycopeptide resistance[END_REF]. Indeed, overexpression or over-activation of σ B induces in some S. aureus strains increased cell wall thickness and promotes increased resistance against teicoplanin, vancomycin and other antibiotics targeting bacterial wall like β-lactams [START_REF] Bischoff | Teicoplanin stress-selected mutations increasing sigma(B) activity in Staphylococcus aureus[END_REF][START_REF] Morikawa | Overexpression of sigma factor, sigma(B), urges Staphylococcus aureus to thicken the cell wall and to resist beta-lactams[END_REF][START_REF] Schulthess | Functional characterization of the sigmaB-dependent yabJ-spoVG operon in Staphylococcus aureus: role in methicillin and glycopeptide resistance[END_REF]). In addition, the phenotypes of a sigB mutant differ depending on the bacterial species, [START_REF] Ferreira | Role of sigma(B) in heat, ethanol, acid, and oxidative stress resistance and during carbon starvation in Listeria monocytogenes[END_REF], Giachino et al. 2001[START_REF] Bandow | Bacillus subtilis tolerance of moderate concentrations of rifampin involves the sigma(B)-dependent general and multiple stress response[END_REF], Zhang et al. 2011, Reder et al. 2012c). The growth and survival of a sigB mutant of B. subtilis or L. monocytogenes is affected in presence of high osmolarity, low pH or high temperature [START_REF] Volker | Expression of the sigmaB-dependent general stress regulon confers multiple stress resistance in Bacillus subtilis[END_REF][START_REF] Ferreira | Role of sigma(B) in heat, ethanol, acid, and oxidative stress resistance and during carbon starvation in Listeria monocytogenes[END_REF][START_REF] Wemekamp-Kamphuis | Identification of sigma factor sigma B-controlled genes and their impact on acid stress, high hydrostatic pressure, and freeze survival in Listeria monocytogenes EGD-e[END_REF]). In agreement, σ B of B. subtilis controls the transcription of genes required for the uptake of solutes (opuD and opuE) mediating osmoprotection [START_REF] Von Blohn | Osmostress response in Bacillus subtilis: characterization of a proline uptake system (OpuE) regulated by high osmolarity and the alternative transcription factor sigma B[END_REF], while in L. monocytogenes, many identified σ B -controlled genes participate in the stress response to acidification (e.g., gadCB, gadD) or exposure to bile salts (e.g., bsh, opuC, bilE). This supports that σ B is involved in multiple protective mechanisms against the challenges faced by L. monocytogenes during gastrointestinal infection and food processing [START_REF] Sue | Sigma(B)-dependent expression patterns of compatible solute transporter genes opuCA and lmo1421 and the conjugated bile salt hydrolase gene bsh in Listeria monocytogenes[END_REF][START_REF] Sleator | A PrfAregulated bile exclusion system (BilE) is a novel virulence factor in Listeria monocytogenes[END_REF]). An ethanol sensitivity for a sigB mutant is observed only in B. subtilis [START_REF] Volker | Expression of the sigmaB-dependent general stress regulon confers multiple stress resistance in Bacillus subtilis[END_REF] while sigB inactivation in B. cereus causes a decreased temperature resistance (van Schaik et al. 2004). σ B is also involved in the oxidative-stress response in B. subtilis, L. monocytogenes and S. aureus. Indeed, the viability of a sigB mutant of L. monocytogenes and S. aureus decreases when exposed to H 2 O 2 or H 2 O 2 -donor compounds [START_REF] Ferreira | Role of sigma(B) in heat, ethanol, acid, and oxidative stress resistance and during carbon starvation in Listeria monocytogenes[END_REF][START_REF] Cebrian | Role of the alternative sigma factor sigma on Staphylococcus aureus resistance to stresses of relevance to food preservation[END_REF]). In B. subtilis, the sigB mutant is more sensitive to oxidative stress when cells are pre-adapted to a stress such as glucose starvation known to induce the activity of σ B before exposure to H 2 O 2 or paraquat (a superoxide donor) (Engelmann andHecker 1996, Reder et al. 2012c). This was consistent with genes positively controlled by σ B in B. subtilis encoding the KatB, KatX and YdbD catalases as well as thioredoxin TrxA, involved in thiol homeostasis [START_REF] Scharf | Thioredoxin is an essential protein induced by multiple stresses in Bacillus subtilis[END_REF][START_REF] Petersohn | Global analysis of the general stress response of Bacillus subtilis[END_REF], Price et al. 2001). Moreover the gene encoding the Dps protein involved in DNA protection is also controlled by σ B [START_REF] Antelmann | Expression of a stress-and starvation-induced dps/pexB-homologous gene is controlled by the alternative sigma factor sigmaB in Bacillus subtilis[END_REF], Wang et al. 2009b). However, the role of σ B in the response to an NO stress stimuli is much less clear. Indeed, in B. subtilis, even if sodium nitroprusside (SNP), a NO donor or NO gas induces the expression of some σ B -controlled genes under aerobic conditions [START_REF] Moore | Response of Bacillus subtilis to nitric oxide and the nitrosating agent sodium nitroprusside[END_REF][START_REF] Rogstam | Mechanisms of adaptation to nitrosative stress in Bacillus subtilis[END_REF]), a sigB mutant is not more sensitive to SNP [START_REF] Rogstam | Mechanisms of adaptation to nitrosative stress in Bacillus subtilis[END_REF]. Finally, in addition to stress response, σ B can regulates the expression of virulence genes. A sigB mutant of L. monocytogenes shows a reduced virulence in a murine model, which suggested a role of σ B in the pathogenesis of this bacterium (Wiedmann et al. 1998[START_REF] Nadon | Sigma B contributes to PrfAmediated virulence in Listeria monocytogenes[END_REF]. In fact, σ B positively controls the expression of genes encoding the internalins InlA and InlB, which are involved in the internalization of this bacterium through interaction with E-cadherin or the Met and gC1qR receptors [START_REF] Mengaud | E-cadherin is the receptor for internalin, a surface protein required for entry of L. monocytogenes into epithelial cells[END_REF][START_REF] Kazmierczak | Listeria monocytogenes sigma B regulates stress response and virulence functions[END_REF], Kim et al. 2005[START_REF] Garner | Sigma B contributes to Listeria monocytogenes gastrointestinal infection but not to systemic spread in the guinea pig infection model[END_REF][START_REF] Mcgann | The alternative sigma factor sigma B and the virulence gene regulator PrfA both regulate transcription of Listeria monocytogenes internalins[END_REF]). In S. aureus, several genes involved in virulence seem to be under σ B control including genes encoding several adhesins [START_REF] Bischoff | Microarray-based analysis of the Staphylococcus aureus sigmaB regulon[END_REF][START_REF] Entenza | Role of sigmaB in the expression of Staphylococcus aureus cell wall adhesins ClfA and FnbA and contribution to infectivity in a rat model of experimental endocarditis[END_REF].

As observed for the signaling pathway, considerable differences exist among species concerning the functions regulated by this alternative σ factor. In Firmicutes, the role of the σ B regulon may have evolved to probably facilitate bacterial survival according with the lifestyle, the niche colonized and the stresses encountered by each species. as for other Firmicutes, and its infectious cycle. σ B positively and negatively controls the expression of about 1000 genes of C.difficile (about ~25% of genes) including genes involved in metabolism, motility, sporulation, cell surface biogenesis and stress management (Figure 11) (Kint et al. 2017). It was noted that 410 genes are positively controlled by σ B corresponding to the largerst number of genes when compared to the other Firmicutes [START_REF] Bischoff | Microarray-based analysis of the Staphylococcus aureus sigmaB regulon[END_REF][START_REF] Nannapaneni | Defining the structure of the general stress regulon of Bacillus subtilis using targeted microarray analysis and random forest classification[END_REF], Guldimann et al. 2016[START_REF] Mader | Staphylococcus aureus Transcriptome Architecture: From Laboratory to Infection-Mimicking Conditions[END_REF]. Kint et al., showed that a sigB mutant of C. difficile is more sensitive to acidification and CAMPs as well as to toxic compounds produced by the immune system such as H 2 O 2 , superoxide anion or NO (Kint et al. 2017). Moreover, it was shown for the first time in this bacterium, that σ B plays a crucial role in O 2 tolerance. In agreement, σ B positively controls genes encoding proteins potentially involved in O 2 , ROS and NO detoxification. This includes CD1157 and CD1623 encoding flavodiiron proteins, two genes (CD1125 and CD0837) encoding nitro-reductases, CD0827 encoding a desulfoferrodoxin, CD1474 and CD1524 encoding reverse-rubrerythrins, and CD0176 encoding a protein similar to NADHoxidoreductase (NADH-OR) supporting an important role of σ B in the management of oxidative and nitrosative stress. This σ factor also controls thiol homeostasis and iron-sulfur clusters (Fe-S) biogenesis. Genes encoding proteins involved in thiol protection (CD1690-a thioredoxin (Trx), and CD1822-a thiol-peroxidase) and biogenesis of Fe-S centers (CD1279 and CD3670, cysteine desulfurases) are also positively controlled by σ B . These functions are important for maintenance of the metabolic activity of anaerobes that uses many Fe-S containing enzymes [START_REF] Meyer | Clostridial iron-sulphur proteins[END_REF] and for counteracting the effects of oxidizing compounds.

II

In the proximal colon, the pH can reach acidic levels due to short-chain fatty-acid production [START_REF] Nugent | Intestinal luminal pH in inflammatory bowel disease: possible determinants and implications for therapy with aminosalicylates and other drugs[END_REF]. The growth of the sigB mutant being severely affected at pH 5.0 suggests that σ B controls, at least partly, genes involved in the acidic stress management and allow the C. difficile vegetative cells to cope with pH variations along the intestinal tract.

However, the molecular mechanisms underlying the adaptation to acidic stress have yet to be elucidated in C. difficile. Additionally, σ B seems to participate in the cell envelope homeostasis by controlling the expression of several genes encoding membrane-and cell Importantly, C. difficile sigB mutant has a significant colonization defect in the axenic mouse when competing with the wild-type strain (Kint et al. 2017). The decreased ability of the sigB mutant to manage stresses encountered during gut colonization is probably responsible in part for the colonization defect. Moreover the control by σ B of the genes encoding cell surfaceassociated proteins, whose some of them are involved in adhesion (Janoir 2016), might explain the failure to detect cells of the sigB mutant associated with the caecal mucosa. These results suggest a contribution of this sigma factor in the ability of C. difficile to adhere to intestinal cells (Kint et al. 2017). In conclusion, σ B has been proposed as a key regulator of gut colonization by coordinating several processes likely required for adaptation, stress management and survival of bacteria within the gut (Kint et al. 2017).

II.4.a. Characterization of the σ B signaling pathway in C. difficile

In C. difficile, sigB (CD0011) is the last gene of a five-gene operon that contains the genes encoding RsbV (CD0009) and RsbW (CD0010) (Figure 12). The two first genes of the operon, CD0007 and CD0008, encode two small proteins of unknown function. RsbV and RsbW of C. difficile share 35% and 39% of identity with RsbV and RsbW of B. subtilis, respectively. In other firmicutes, genes near sigB gene usually encode proteins involved in the signaling pathway leading to the activation of σ B (Hecker et al. 2007). However, in C.

difficile, such proteins involved in the activation of σ B are still unknown. I have participated in the study of the signaling pathway leading to the activation of σ B in C. difficile in collaboration with Nicolas Kint (Kint et al., in preparation; see Article in Annex). In this study, my main contribution was on the analysis of the expression of the sigB gene and its activity during vegetative growth, using transcriptional fusions and fluorescence microscopy (Kint et al, in Annex).

2017), confirming the involvement of these two proteins in σ B activation. As in other Firmicutes, σ B is kept inactive by RsbW (Figure 9B). In the presence of a stress signal, RsbV is dephosphorylated, binds to RsbW leading to the release of σ B and the transcription of its targets. No obvious role for CD0007 and CD0008 was observed in σ B activation under the tested conditions. The inactivation of CD0007 and CD0008 genes had no impact on the sensitivity of C. difficile to low O 2 tension or NO-donor compounds and did not modify the expression of σ B -target genes at the onset of the stationary phase.

Players and signals in the activation of σ B

The dephosphorylation of RsbV leading to the activation of σ B is usually mediated by specific PP2C phosphatase(s) [START_REF] Louie | OPT-80 eliminates Clostridium difficile and is sparing of bacteroides species during treatment of C. difficile infection[END_REF][START_REF] Tannock | A new macrocyclic antibiotic, fidaxomicin (OPT-80), causes less alteration to the bowel microbiota of Clostridium difficile-infected patients than does vancomycin[END_REF][START_REF] Petrella | Decreased cure and increased recurrence rates for Clostridium difficile infection caused by the epidemic C. difficile BI strain[END_REF] tensions as the sigB mutant (Kint et al. 2017). Moreover, the expression of σ B -target genes decreased when the rsbZ gene was inactivated. In addition to its phosphatase domain, RsbZ has a GAF domain. The GAF domain of RsbZ might be involved in the detection of nutritional stresses in C. difficile. 13). Some obligate anaerobes contain SOD and/or catalase, but the latter is less distributed among anaerobes than SOD [START_REF] Brioukhanov | Catalase and superoxide dismutase: distribution, properties, and physiological role in cells of strict anaerobes[END_REF]. The sulfate-reducing bacteria Desulfovibrio gigas contains both a SOD and a catalase, even if catalase activity does not seem to be present in all Desulfovibrio species (Dos [START_REF] Santos | Purification and characterization of an iron superoxide dismutase and a catalase from the sulfate-reducing bacterium Desulfovibrio gigas[END_REF]. Desulfovibrio vulgaris and D. gigas are catalase positive, and for the latter one long exposure times to O 2 lead to higher catalase activity [START_REF] Fareleira | Response of a strict anaerobe to oxygen: survival strategies in Desulfovibrio gigas[END_REF]. The obligate anaerobic gingival pathogenic Porphyromonas gingivalis also contains a SOD but lacks a catalase (Amano et al. 1986, Lynch and[START_REF] Lynch | Role of superoxide dismutase activity in the physiology of Porphyromonas gingivalis[END_REF]. In the genome of the strict anaerobe C. acetobutylicum, no catalase was annotated, and no function has been yet demonstrated for the annotated SODs [START_REF] Nolling | Genome sequence and comparative analysis of the solvent-producing bacterium Clostridium acetobutylicum[END_REF]. [START_REF] Briolat | Identification of the Clostridium perfringens genes involved in the adaptive response to oxidative stress[END_REF]. This supported that alternative detoxification systems must exist. Indeed, anaerobic and microaerophilic bacteria use specific reduction pathways for ROS detoxification based

I.2. Alternative detoxification systems: NADH-dependent reductases

revRbr N C

mainly on NADH-dependent reductases that lead to water rather than O 2 production [START_REF] Jenney | Anaerobic microbes: oxygen detoxification without superoxide dismutase[END_REF][START_REF] Lumppio | Rubrerythrin and rubredoxin oxidoreductase in Desulfovibrio vulgaris: a novel oxidative stress protection system[END_REF], Hillmann et al. 2009b, Riebe et al. 2009). These organisms use superoxide reductases (SORs), rubrerythrins (Rbrs), reverse rubrerythrins (revRbrs) and flavodiiron proteins (FPDs) (Figure 13). These enzymes have been isolated and characterized from a number of organisms, including D. vulgaris [START_REF] Wildschut | Rubredoxin:oxygen oxidoreductase enhances survival of Desulfovibrio vulgaris hildenborough under microaerophilic conditions[END_REF], P. gingivalis [START_REF] Lynch | Role of superoxide dismutase activity in the physiology of Porphyromonas gingivalis[END_REF]Kuramitsu 1999, Sztukowska et al. 2002), C. perfringens [START_REF] Jean | Oxidative stress response in Clostridium perfringens[END_REF]) and C.

acetobutylicum [START_REF] Riebe | Desulfoferrodoxin of Clostridium acetobutylicum functions as a superoxide reductase[END_REF]

I.2.a. Rubrerythrins (Rbrs)

Rubrerythrins (Rbrs) constitute a family of proteins that are implicated in oxidative stress protection in anaerobic bacteria and archaea. These proteins contain two types of iron sites: an N-terminal non-sulfur diiron center and a rubredoxin-type [FeCys 4 ] center, located at the C-terminal part (Figure 14) [START_REF] Demare | The structure of Desulfovibrio vulgaris rubrerythrin reveals a unique combination of rubredoxin-like FeS4 and ferritin-like diiron domains[END_REF]. Reverse Rbr (revRbr) are enzymes in which the position of the diiron site and the rubredoxin domains are reversed (Figure 14).

Both Rbr and revRbr have been shown to play a role as NAD(P)H:H 2 O 2 oxidoreductase [START_REF] Coulter | NADH peroxidase activity of rubrerythrin[END_REF], Weinberg et al. 2004). The reduced diiron site reacts directly with H 2 O 2 , while the [Fe(SCys) 4 ] site transfers electrons from an exogenous donor to the oxidized diiron site. NAD(P)H-dependent peroxidase activity was shown for classical Rbrs from D. vulgaris [START_REF] Lumppio | Rubrerythrin and rubredoxin oxidoreductase in Desulfovibrio vulgaris: a novel oxidative stress protection system[END_REF] and Pyrococcus furiosus [START_REF] Weinberg | Rubrerythrin from the hyperthermophilic archaeon Pyrococcus furiosus is a rubredoxin-dependent, iron-containing peroxidase[END_REF]). An O 2 -reductase activity was also described for these proteins. However, the reactivity of this enzymes to O 2 is typically low relative to that with H 2 O 2 . In vitro, the D. vulgaris Rbr reduces H 2 O 2 much more rapidly than O 2 [START_REF] Coulter | NADH peroxidase activity of rubrerythrin[END_REF]Kurtz 2001). An increased sensitivity to both O 2 and H 2 O 2 was observed in Rbr-deletion mutant of P. gingivalis [START_REF] Sztukowska | Role of rubrerythrin in the oxidative stress response of Porphyromonas gingivalis[END_REF].

C. acetobutylicum genome encodes four proteins homologous to Rbrs: 2 normal Rbrs (RubY and RubZ) and 2 revRbs (Rbr3B and Rbr3A whose encoding genes forms an operon) [START_REF] May | A rubrerythrin-like oxidative stress protein of Clostridium acetobutylicum is encoded by a duplicated gene and identical to the heat shock protein Hsp21[END_REF]. While normal Rbrs are upregulated upon oxygenation, revRbrs seems to be induced both upon oxygenation and oxidative stress [START_REF] Hillmann | The rubrerythrin-like protein Hsp21 of Clostridium acetobutylicum is a general stress protein[END_REF]). RubY has a peroxidase specific activity comparable to that of the revRbr in vitro, [START_REF] May | A rubrerythrin-like oxidative stress protein of Clostridium acetobutylicum is encoded by a duplicated gene and identical to the heat shock protein Hsp21[END_REF][START_REF] Hillmann | The rubrerythrin-like protein Hsp21 of Clostridium acetobutylicum is a general stress protein[END_REF], Riebe et al. 2009), while the revRbrs, that are 99% identical, have both NADH peroxidase and oxidase activity [START_REF] Kawasaki | Identification of O2-induced peptides in an obligatory anaerobe, Clostridium acetobutylicum[END_REF][START_REF] May | A rubrerythrin-like oxidative stress protein of Clostridium acetobutylicum is encoded by a duplicated gene and identical to the heat shock protein Hsp21[END_REF], Riebe et al. 2009).

The overproduction of the Rbr3B showed significantly increased tolerance to O 2 and H 2 O 2 exposure [START_REF] Riebe | Pathway for H2O2 and O2 detoxification in Clostridium acetobutylicum[END_REF]). H 2 O 2 and O 2 detoxification pathway in C. acetobutylicum involves a combination of several proteins i.e., Rbr or revRbr, rubredoxin (Rd) and NADHrubredoxin oxidoreductase (NROR) that use NADH as an electron donor [START_REF] Kawasaki | O2 and reactive oxygen species detoxification complex, composed of O2-responsive NADH:rubredoxin oxidoreductase-flavoprotein A2-desulfoferrodoxin operon enzymes, rubperoxin, and rubredoxin, in Clostridium acetobutylicum[END_REF], Riebe et al. 2009). Rbrs/revRbrs eliminate H 2 O 2 and/or O 2 through their reduction to water [START_REF] Kawasaki | Identification of O2-induced peptides in an obligatory anaerobe, Clostridium acetobutylicum[END_REF][START_REF] May | A rubrerythrin-like oxidative stress protein of Clostridium acetobutylicum is encoded by a duplicated gene and identical to the heat shock protein Hsp21[END_REF][START_REF] Hillmann | The rubrerythrin-like protein Hsp21 of Clostridium acetobutylicum is a general stress protein[END_REF] [START_REF] Hillmann | PerR acts as a switch for oxygen tolerance in the strict anaerobe Clostridium acetobutylicum[END_REF], Hillmann et al. 2009a). In fact, the inactivation of perR is related to an increased tolerance to O 2 and higher resistance to H 2 O 2, leads to an increased expression of genes coding for proteins involved in the detoxification of O 2 and H 2 O 2 [START_REF] Hillmann | PerR acts as a switch for oxygen tolerance in the strict anaerobe Clostridium acetobutylicum[END_REF].

I.2.b. Flavodiiron proteins (FDPs)

The flavodiiron proteins, called FDPs, are also involved in O 2 scavenging. FDPs have O 2 and/or NO reductase activity and belong to a family widespread in the three domains of life. These enzymes are found both in aerobic and anaerobic organisms. Based on their different domains, several classes of FDPs from A to H have been proposed (Figure 15) (Folgosa et al. 2018a). Class A FDPs contains only the minimal core composed of two domains: a metallo-β-lactamase-like domain, containing a diiron catalytic center, and a flavodoxin harboring a flavin mononucleotide (FMN). FDP activity requires also the participation of proteins involved in electron transfer such as Rds and NADH/FAD dependent oxidoreductases [START_REF] Chen | Rubredoxin oxidase, a new flavo-hemo-protein, is the site of oxygen reduction to water by the "strict anaerobe" Desulfovibrio gigas[END_REF][START_REF] Silaghi-Dumitrescu | A flavodiiron protein and high molecular weight rubredoxin from Moorella thermoacetica with nitric oxide reductase activity[END_REF] or F 420 coenzyme in methanogens (Figure 16) [START_REF] Seedorf | F420H2 oxidase (FprA) from Methanobrevibacter arboriphilus, a coenzyme F420-dependent enzyme involved in O2 detoxification[END_REF]). In some cases (see below), the Rd or the NADH/FAD dependent oxidoreductase domain are fused to the FDP core domain or present as independent proteins (Figure 15 and 16). Class A FDPs have either a clear preference for O 2 as observed for protozoal and archaeal enzymes [START_REF] Seedorf | F420H2 oxidase (FprA) from Methanobrevibacter arboriphilus, a coenzyme F420-dependent enzyme involved in O2 detoxification[END_REF][START_REF] Smutna | Flavodiiron protein from Trichomonas vaginalis hydrogenosomes: the terminal oxygen reductase[END_REF][START_REF] Vicente | A detoxifying oxygen reductase in the anaerobic protozoan Entamoeba histolytica[END_REF] or exhibit a dual activity towards NO and O 2 as described for the enzymes of M. thermoacetica or D. gigas [START_REF] Silaghi-Dumitrescu | A flavodiiron protein and high molecular weight rubredoxin from Moorella thermoacetica with nitric oxide reductase activity[END_REF][START_REF] Rodrigues | Desulfovibrio gigas flavodiiron protein affords protection against nitrosative stress in vivo[END_REF]. C. acetobutylicum possesses two FDPs, FprA1 and FprA2, with NADH:O 2 oxidoreductase activity that use NROR and a Rd as electron transporters (Hillmann et al. 2009b). FprA1 and FprA2 are more produced in the perR mutant strain [START_REF] May | A rubrerythrin-like oxidative stress protein of Clostridium acetobutylicum is encoded by a duplicated gene and identical to the heat shock protein Hsp21[END_REF]) and their encoding genes are induced after O 2 exposure or during oxidative stress [START_REF] Kawasaki | Identification of O2-induced peptides in an obligatory anaerobe, Clostridium acetobutylicum[END_REF][START_REF] Kawasaki | Adaptive responses to oxygen stress in obligatory anaerobes Clostridium acetobutylicum and Clostridium aminovalericum[END_REF]. Those proteins, together with the Rbrs and Dfx form an O 2 and H 2 O 2 detoxification complex in C. acetobutylicum that is schematically represented in the Figure 17. The Class B FDPs, which includes NorV from Escherichia coli, is restricted to the Proteobacteria phylum. In addition to the core domain, NorV contains a Rd domain with a FeCys4 center and for this reason was called flavorubredoxin (Figure 16) [START_REF] Gomes | Spectroscopic studies and characterization of a novel electron-transfer chain from Escherichia coli involving a flavorubredoxin and its flavoprotein reductase partner[END_REF], Gomes et al. 2002). This protein receives electrons from an NROR called NorW. E. coli NorV is the only example so far of an FDP with a clear preference for NO as a substrate (Vicente andTeixeira 2005, Vicente et al. 2007). It was shown that the transcriptional levels of the norV gene were higher after exposing E .coli cells to NO and NorV confers resistance to NO [START_REF] Gardner | Flavorubredoxin, an inducible catalyst for nitric oxide reduction and detoxification in Escherichia coli[END_REF][START_REF] Justino | New genes implicated in the protection of anaerobically grown Escherichia coli against nitric oxide[END_REF]. Finally, the Class C FDPs is widely distributed in cyanobacteria and contains a C-terminal flavin-binding domain. Class C FDP was isolated from Synechocystis sp and was shown to have NADH: O 2 oxidoreductase activity [START_REF] Wasserfallen | A family of flavoproteins in the domains Archaea and Bacteria[END_REF], Vicente et al. 2002). Class F FDPs occur mainly in Firmicutes, namely from the Clostridiales order and one example was recently characterized in C. difficile (see more in detail Chapter 3, section II. O 2 and ROS detoxification in C. difficile). For the remaining four classes of FDPs, their properties are deduced from the respective amino acid sequences, due to the lack of reported studies.

II. O 2 and ROS detoxification in C. difficile

II.1. The exposure of C. difficile to O 2 , ROS and NO during infection

During its infectious cycle, C. difficile vegetative cells are gradually exposed to O 2 and oxidative stress. Indeed, the gut is not strictly anaerobic (Espey 2013) and an increasing gradient of O 2 exist from the intestinal lumen (anaerobic) to the mucosa (approximately 5%

O 2 ). After an antibiotic treatment, the gradient of O 2 can be disturbed and cause an increase of O 2 tension in the gut (Rivera-Chavez et al. 2017). Indeed, the composition of gut-associated microbial communities is modified during antibiotherapy. For instance, a depletion of butyrate-producing microorganisms limits the production of butyrate and its oxidation by epithelial cells increasing diffusion of O 2 into the intestinal lumen (Figure 18) [START_REF] Rivera-Chavez | Depletion of Butyrate-Producing Clostridia from the Gut Microbiota Drives an Aerobic Luminal Expansion of Salmonella[END_REF], Rivera-Chavez et al. 2017). Moreover, the inflammatory response triggered by C.

difficile is a source of oxidative stress. When recruited to the infection site, immune cells such as macrophages and neutrophils ingest the invading bacteria and bombard them with oxidants. release of iron that lead to oxidative stress through the fenton reaction [START_REF] Imlay | Iron-sulphur clusters and the problem with oxygen[END_REF].

Moreover, H 2 O 2 , which is able to penetrates lipid bilayers similar to that of water [START_REF] Seaver | Hydrogen peroxide fluxes and compartmentalization inside growing Escherichia coli[END_REF], can oxidize lipids, cysteine residues of proteins and Fe-S clusters [START_REF] Imlay | Pathways of oxidative damage[END_REF][START_REF] Semchyshyn | Hydrogen peroxide increases the activities of soxRS regulon enzymes and the levels of oxidized proteins and lipids in Escherichia coli[END_REF]. Finally, NO can modify cysteine residues (S-nitrosylation) and react with transition metals (Vazquez-Torres and Baumler 2016), while ONOO -oxidizes thiols, DNA and the transition metal centers of metalloproteins [START_REF] Radi | Peroxynitrite, a stealthy biological oxidant[END_REF].

II.2. The response of C. difficile to O 2 and oxidative stress

The response of C. difficile to O 2 , oxidative and nitrosative stresses is poorly studied.

Vegetative cells of this strictly anaerobic bacterium, are known to be highly sensitive to low levels of O 2 [START_REF] Holy | Oxygen tolerance in anaerobic pathogenic bacteria[END_REF]. However, it has been recently demonstrated that this pathogen is able to grow in non-strict anaerobic conditions and to adapt to low O 2 tension.

Indeed, this bacterium is able to grow under 2% O 2 (Giordano et al. 2018a) and 5% O 2 has only a moderate effect on C. difficile growth [START_REF] Neumann-Schaal | Tracking gene expression and oxidative damage of O2stressed Clostridioides difficile by a multi-omics approach[END_REF]. However, the tolerance to O 2 seems to be strain-dependent. The survival of the 630Δerm reference strain is significantly reduced only after 3 hours of exposure to air and 12 hours are necessary to completely eliminate all vegetative cells. The "highly virulent" strain R20291 of 027 ribotype, as well as a clinical strain of 078 ribotype are less O 2 -tolerant with a survival that decreased sharply after 1 hour of exposure to air and no bacteria quantified after 3 hours of air exposure (Edwards et al. 2016a). In addition, C. difficile vegetative cells are sensitive to relative low concentrations of H 2 O 2 (400 µM). As observed with air, the sensitivity to H 2 O 2 exposure is also strain-dependent. The 630Δerm strain is more resistant than strains of the 027 and 078 ribotypes (Edwards et al. 2016a [START_REF] Girinathan | Clostridium difficile glutamate dehydrogenase is a secreted enzyme that confers resistance to H2O2[END_REF]. This effect might be due to a decreased concentration of α-ketoglutarate, a compound that might be able of capturing H 2 O 2 [START_REF] Mailloux | Alphaketoglutarate dehydrogenase and glutamate dehydrogenase work in tandem to modulate the antioxidant alpha-ketoglutarate during oxidative stress in Pseudomonas fluorescens[END_REF]. The deletion of the cysteine desulfurase encoding gene iscS involved in the Fe-S cluster (Isc) system cause a severe growth defect of C. difficile in the presence of 2% O 2 and the mutant fail to colonize a germ-free model (Giordano et al. 2018b [START_REF] Emerson | Microarray analysis of the transcriptional responses of Clostridium difficile to environmental and antibiotic stress[END_REF]. They found an increased expression of an operon encoding a reductase (CD0174-76) and of genes encoding a thiol peroxidase (CD1822), a Trx-Trx reductase system (CD1690-91) and a revRbr (CD1524). The high homology (96% identity) of the two genes encoding the revRbrs (CD1474 and CD1524) did not allow determining if only one or both are upregulated.

Expression of the two genes encoding the FDPs (CD1623 and CD1157) probably involved in O 2 and/or NO reduction also increased in the presence of air [START_REF] Emerson | Microarray analysis of the transcriptional responses of Clostridium difficile to environmental and antibiotic stress[END_REF]. However, in two more recent transcriptomic or multi-omic studies, no induction of these genes was observed under microaerophillic growth conditions (Giordano et al. 2018a[START_REF] Neumann-Schaal | Tracking gene expression and oxidative damage of O2stressed Clostridioides difficile by a multi-omics approach[END_REF]. In those studies, the greatest expression changes concerned genes encoding proteins involved in sugar and amino acid metabolism. Changes in the reductive pathways of the Stickland reaction for the use of proline, glycine or leucine were observed. In addition, an induction of genes for the de-novo synthesis of cysteine was verified, likely associated with an important oxidation of cysteine in proteins in microaerophillic growth conditions. Carbon metabolism was also significantly modified with changes in the phosphotransferase system (PTS) facilitating uptake of sugars and a repression of the synthesis of enzymes involved in glycolysis and butyrate fermentation. A temporary induction in the synthesis of cofactor riboflavin was also detected possibly due to a demand for FMN and FAD in redox reactions and CotG-CD1567). However, the genes encoding these proteins are only expressed during sporulation and likely transcribed by the late sporulation sigma factors (σ G and σ K ) (Fimlaid et al. 2013, Permpoonpattana et al. 2013, Saujet et al. 2013). Moreover, those proteins are located at the surface of C. difficile spore (Lawley et al. 2009b, Abhyankar et al. 2013, Diaz-Gonzalez et al. 2015).

Nevertheless Moreover, σ B and σ H inversely control the sporulation process (Saujet et al. 2011, Kint et al. 2017). σ B negatively controls the expression of more than 200 genes transcribed by the different sporulation-specific σ factors. In agreement, a sigB mutant sporulates more efficiently and/or earlier than the wild-type strain (Kint et al. 2017). By contrast, σ H positively controls several genes required for sporulation and, accordingly, sigH inactivation results in an asporogeneous phenotype (Saujet et al. 2011). During transition from exponential to stationary growth phase, cells will activate a complex regulatory network to promote the more suitable choice to make in response to the given conditions. Thus, under stress conditions, such as entering stationary phase, C. difficile must quickly choose between a rapid adaptative response mediated by σ B , or an irreversible and slower differentiation process leading to the formation of spores whose initiation is controlled by σ H . Sporulation is predominantly the last resource for cells experiencing nutrient limitation.

II. The sporulation process

Endospores are formed by three classes of low G+C Gram positive bacteria (bacilli, clostridia and negativicutes), as a strategy to survive to unfavorable conditions. Bacterial

σ F σ E σ F σ E σ K σ G σ K σ G σ K σ G
spores resist to extremes of physical and chemical parameters that would rapidly destroy vegetative cells, being the most resistant cellular structure known [START_REF] Setlow | I will survive: DNA protection in bacterial spores[END_REF]. Spore forming bacteria include important organisms in several fields. Certain species such as Clostridium acetobutylicum, a saprophyte clostridium able to undertake useful biotransformations while others are important animal and human pathogens responsible for diseases such as anthrax, botulism, diarrhea, gas gangrene or tetanus [START_REF] Al-Hinai | The Clostridium sporulation programs: diversity and preservation of endospore differentiation[END_REF].

For pathogenic bacteria, sporulation is crucial to survive during transmission from one host to another. Sporulation is a process of cellular differentiation allowing the conversion of a vegetative cell into a mature spore through several morphological stages (Figure 19). The main morphological stages are conserved between species (Piggot andCoote 1976, Fimlaid andShen 2015). Stage 0 corresponds to the stage where the cells are undergoing normal vegetative growth by binary fission. When the growth conditions are unfavorable, the bacteria enter Stage I during which the chromosome is replicated and segregated in a polar way. Then, sporulation starts when an asymmetric division occurs near to one pole of the vegetative cell, resulting in the formation of a smaller cell, the forespore (FS) and a larger cell, the mother cell (MC). Each compartment contains a copy of the chromosome (Stage II). Evidences have indicated that the division can take place either in the "old" or the "new" cell pole [START_REF] Veening | A mechanism for cell cycle regulation of sporulation initiation in Bacillus subtilis[END_REF]). The sporulation septum contains a thinner layer of peptidoglycan (PG) separating the two compartments. After asymmetric division the MC engulfs the FS in a phagocytic-like manner (Stage III). During stage IV, the spore cortex is formed between the two membranes of the MC and the FC. Stage V corresponds to the assembly of the spore coat and stage VI is the maturation of the spore layers, cortex and coat. At the end (Stage VII), the mature spore is released to the environment through lysis of the MC (Hilbert and Piggot 2004). Although sporulation takes about 8-10 hours, vegetative growth is resumed in only a few minutes, as soon as the spore senses its germinants, following a process called germination (see more in detail Chapter 4, section III. Germination).

II.1. Regulation of sporulation in B. subtilis

Sporulation initiation is controlled by the master regulator Spo0A, present in all spore formers (Paredes et al. 2005[START_REF] De Hoon | Hierarchical evolution of the bacterial sporulation network[END_REF], Galperin et al. 2012, Abecasis et al. 2013).

In bacilli and clostridia, the development program of sporulation is mainly governed by four cell-type specific sigma factors: σ F , σ E , σ G or σ K . In B. subtilis, the first sigma factors, σ F in the FS and σ E in the MC, are activated soon after asymmetric division and control early stages Spo0A phosphorylation is controlled by a phosphorelay that depends on the activity of several kinases and other intermediaries. Interactions that ultimately lead to Spo0A phosphorylation are indicated in black, while the ones working against it are in red. The KinA and KinB kinases respond to yet not known signals by autophosphorylating and subsequently transferring phosphate to Spo0F. Spo0F phosphorylates Spo0B, which passes phosphate on to Spo0A. A third kinase, KinC, bypasses the phosphorelay and acts directly on Spo0A. Phosphatases, including RapA, B and E and Spo0E, and kinase inhibitors (Sda) act at various steps in the pathway to directly or indirectly antagonize Spo0A phosphorylation.

of spore development (Figure 19). σ G and σ K replace σ F and σ E in their respective compartments and control gene expression following engulfment completion (Figure 19) [START_REF] Stragier | Molecular genetics of sporulation in Bacillus subtilis[END_REF], Hilbert and Piggot 2004, Higgins and Dworkin 2012). Therefore, inactivation of both sigF and sigE blocks the sporulation process soon after asymmetric division generating cells presenting a disporic phenotype, while inactivation of sigG and sigK results in a blockage of the process after engulfment completion (Piggot andCoote 1976, Stragier andLosick 1996). The activation of σ G and σ K requires the preceding σ F and σ E factors to be active in the respective cell compartment [START_REF] Stragier | Molecular genetics of sporulation in Bacillus subtilis[END_REF][START_REF] Errington | Regulation of endospore formation in Bacillus subtilis[END_REF], Hilbert and Piggot 2004, Higgins and Dworkin 2012). Thus, after asymmetric division two parallel programs of gene expression define the FS and the MC lines of gene expresion.

In B. subtilis, six transcriptional regulators (SpoIIID, GerR, GerE, SpoVT, RfsA and YlyA) work in association with the different σ factors to modulate the expression of their targets (Figure 20) (de Hoon et al. 2010, Higgins andDworkin 2012).

II.1.a Initiation of sporulation

Nutrient starvation has been pointed as the triggering factor to start the sporulation process in many endospore formers, notably in B. subtilis and other bacilli [START_REF] Stephens | Bacterial sporulation: a question of commitment?[END_REF].

Previous studies revealed that the nutritional sensor protein, CodY, negatively regulates sporulation [START_REF] Ratnayake-Lecamwasam | Bacillus subtilis CodY represses early-stationary-phase genes by sensing GTP levels[END_REF][START_REF] Molle | Additional targets of the Bacillus subtilis global regulator CodY identified by chromatin immunoprecipitation and genome-wide transcript analysis[END_REF], Piggot and Hilbert 2004) by controling genes directly involved in sporulation initiation, including spo0A, rapA, rapC, rapE, sinIR, sigH and kinB genes (Ratnayake- [START_REF] Ratnayake-Lecamwasam | Bacillus subtilis CodY represses early-stationary-phase genes by sensing GTP levels[END_REF][START_REF] Molle | Additional targets of the Bacillus subtilis global regulator CodY identified by chromatin immunoprecipitation and genome-wide transcript analysis[END_REF][START_REF] Chateau | Identification of CodY targets in Bacillus anthracis by genome-wide in vitro binding analysis[END_REF]. The spo0A gene is transcribed from both σ A -and σ H -dependent promoters. Then, phosphorylation of Spo0A (Spo0A-P), leading to its dimerization, dramatically increases its affinity to its target sites on the DNA [START_REF] Lewis | Dimer formation and transcription activation in the sporulation response regulator Spo0A[END_REF]. The unphosphorylated form of Spo0A is inactive but phenotypic differentiation processes can come from the extent of Spo0A phosphorylation and abundance. Spo0A phosphorylated at low level induces biofilm development and nutrient scavenging (i.e. cannibalism) [START_REF] Hamon | The sporulation transcription factor Spo0A is required for biofilm development in Bacillus subtilis[END_REF]Lazazzera 2001, Gonzalez-Pastor et al. 2003). By opposition, higher levels of Spo0A-P induce sporulation (Fujita et al. 2005). In B. subtilis, Spo0A phosphorylation is mediated by a phosphorelay that depends on the activity of five sensor histidine kinases (HK) KinA, KinB, KinC, KinD and KinE (Figure 20) (Perego 2001, Higgins andDworkin 2012). KinA and KinB respond to a signal by autophosphorylating and subsequently phosphorylates Spo0F.

Spo0F transfer the phosphate to Spo0B, which in turn transfers it to Spo0A that can activate transcription of sporulation genes (Figure 20). KinA is the major kinase responsible for (Fujita and Losick 2005). It has three PAS domains that in other proteins sense changes in O 2 levels and redox potential. However, the specific physiological signal(s) that stimulate KinA remain unclear. KinC uses its PAS sensory domais to respond to small-molecule-directed potassium leakage and is able bypass the phosphorelay, to phosphorylate directly Spo0A [START_REF] Lopez | Structurally diverse natural products that cause potassium leakage trigger multicellularity in Bacillus subtilis[END_REF]) (Higgins and Dworkin 2012). KinD was shown to delay the onset of sporulation. How KinD affects Spo0A it is not known but it was hyphothesized that KinD functions as a phosphatase [START_REF] Aguilar | KinD is a checkpoint protein linking spore formation to extracellular-matrix production in Bacillus subtilis biofilms[END_REF]. Kinase inhibitors (KipI and Sda), phosphatases (Spo0E, YnzD and YisI) and Rap proteins act at various steps of the pathway and can directly or indirectly influence the Spo0A phosphorylation (Figure 20). The Sda protein regulates KinA activity by binding to it and preventing its autophosphorylation [START_REF] Rowland | Structure and mechanism of action of Sda, an inhibitor of the histidine kinases that regulate initiation of sporulation in Bacillus subtilis[END_REF]). Damage to DNA or block in DNA replication induce sda expression, and consequently Sda prevents activation of Spo0A by the phosphorelay [START_REF] Burkholder | Replication initiation proteins regulate a developmental checkpoint in Bacillus subtilis[END_REF]. The phosphatase Spo0E directly dephosphorylates Spo0A. The Rap proteins A, B and E, dephosphorylate Spo0F-P and thereby reduce the level of Spo0A~P [START_REF] Smits | Temporal separation of distinct differentiation pathways by a dual specificity Rap-Phr system in Bacillus subtilis[END_REF]). Initially, the Rap proteins were thought to function as phosphatases, but evidences suggest that these proteins may cause dephosphorylation of Spo0F-P by binding to it and stimulating its autophosphatase activity, rather than by functioning directly as phosphatases.

II.1.b. The forespore line of gene expression: σ F and σ G When Spo0A-P levels reaches a threshold, Spo0A-P activates sporulation genes such as the spoIIAA-spoIIAB-spoIIAC operon and the spoIIE gene. spoIIAC encodes σ F , the first sporulation-specific sigma factor to be activated in the FS compartment. The expression of the spoIIA operon is controlled by both Spo0A-P and σ H (Figure 21) [START_REF] Wu | Regulation of transcription of the Bacillus subtilis spoIIA locus[END_REF][START_REF] Burbulys | Initiation of sporulation in B. subtilis is controlled by a multicomponent phosphorelay[END_REF]. Although produced in the predivisional cell, σ F becomes active only after the formation of the asymmetric septum [START_REF] Gholamhoseinian | Timing of spoII gene expression relative to septum formation during sporulation of Bacillus subtilis[END_REF]). In the predivisional cell, the anti-sigma factor SpoIIAB that binds to σ F , phosphorylates the anti-anti-sigma factor SpoIIAA and held σ F inactive. SpoIIE, a serine phosphatase localized to the asymmetric septum, dephosphorylates SpoIIAA-P. Then, SpoIIAA will bind to SpoIIAB releasing an active σ F after completion of asymmetric cell division (Figure 21) [START_REF] Barak | SpoIIE mutants of Bacillus subtilis comprise two distinct phenotypic classes consistent with a dual functional role for the SpoIIE protein[END_REF][START_REF] Feucht | Bifunctional protein required for asymmetric cell division and cell-specific transcription in Bacillus subtilis[END_REF], Hilbert and Piggot 2004). Several mechanisms have been proposed

to explain the preferential dephosphorylation of SpoIIAA-P in the FS compartment. The larger surface area-to-volume ratio of the FS compartment allows dephosphorylated SpoIIAA to reach up a critical level to effectively bind all of SpoIIAB, thus activating σ F [START_REF] Iber | The mechanism of cell differentiation in Bacillus subtilis[END_REF]. Also, a preferential localization of SpoIIE to the FS face of the septum has been pointed for the key activation of σ F [START_REF] Guberman | PSICIC: noise and asymmetry in bacterial division revealed by computational image analysis at sub-pixel resolution[END_REF]. The other mechanism is related with transient chromosome asymmetry of spoIIAB gene, and with the relative instability of SpoIIAB protein. Upon asymmetric cell division, only the origin-proximal one third of the chromosome is trapped into the FS compartment [START_REF] Wu | Prespore-specific gene expression in Bacillus subtilis is driven by sequestration of SpoIIE phosphatase to the prespore side of the asymmetric septum[END_REF]. The spoIIAB gene is almost opposite to the oriC region on the chromosome leading to its transient exclusion from the FS compartment. This, in combination with the proteolysis of the SpoIIAB protein reduces the level of anti-σ F relative to σ F [START_REF] Dworkin | Transient genetic asymmetry and cell fate in a bacterium[END_REF]. σ F transcribes around 50 genes [START_REF] Steil | Genome-wide analysis of temporally regulated and compartment-specific gene expression in sporulating cells of Bacillus subtilis[END_REF], Wang et al. 2006). Among these are i) gpr encoding a protease important for the degradation of small acid soluble proteins (SASPs) during germination [START_REF] Sussman | Cloning, nucleotide sequence, and regulation of the Bacillus subtilis gpr gene, which codes for the protease that initiates degradation of small, acid-soluble proteins during spore germination[END_REF], ii) spoIIR encoding a protein required for the activation of σ E (Figure 21) [START_REF] Hofmeister | Extracellular signal protein triggering the proteolytic activation of a developmental transcription factor in B. subtilis[END_REF][START_REF] Karow | Identification of a gene, spoIIR, that links the activation of sigma E to the transcriptional activity of sigma F during sporulation in Bacillus subtilis[END_REF], iii) spoIIQ encoding a protein required for engulfment and also for the activation of σ G (Figure 21) [START_REF] Broder | Forespore engulfment mediated by a ratchet-like mechanism[END_REF]) and iv)

rsfA encoding the RsfA regulator that activates some σ F -transcribed genes and represses others (such as spoIIR) (Wu andErrington 2000, Wang et al. 2006). σ F also transcribes the expression of spoIIIG, encoding σ G , which replaces σ F in the FS (Figure 21) [START_REF] Sun | Control of transcription of the Bacillus subtilis spoIIIG gene, which codes for the forespore-specific transcription factor sigma G[END_REF], Wang et al. 2006). In B. subtilis, spoIIIG (sigG) gene is located downstream of the spoIIGA-spoIIGB (sigE) operon (Figure 21) [START_REF] De Hoon | Hierarchical evolution of the bacterial sporulation network[END_REF]. Transcription of sigG was thought to be delayed in comparison with the expression of other σ F -controlled genes. However, a very recent study reported that sigG is transcribed by σ F but at very low levels as a consequence of several intrinsic features on the sigG regulatory and coding sequence that dampen its transcription and translation [START_REF] Mearls | Transcription and translation of the sigG gene is tuned for proper execution of the switch from early to late gene expression in the developing Bacillus subtilis spore[END_REF]. Accordingly, a model suggests that sigG is first transcribed by σ F , but the majority of sigG transcription occurs later under the control of σ G itself [START_REF] Mearls | Transcription and translation of the sigG gene is tuned for proper execution of the switch from early to late gene expression in the developing Bacillus subtilis spore[END_REF]. σ G can be detected in the FS towards the end of the engulfment, but it is active only after engulfment completion. Activation of σ G depends on the formation of a channel connecting the MC and the FS compartment (Figure 21). This channel is composed of components encoded by genes transcribed by both σ F and σ E [START_REF] Illing | The spoIIIA locus is not a major determinant of prespore-specific gene expression during sporulation in Bacillus subtilis[END_REF][START_REF] Camp | A novel pathway of intercellular signalling in Bacillus subtilis involves a protein with similarity to a component of type III secretion channels[END_REF][START_REF] Mcfee | Clostridium difficile: emerging public health threat and other nosocomial or hospital acquired infections[END_REF][START_REF] Doan | Novel secretion apparatus maintains spore integrity and developmental gene expression in Bacillus subtilis[END_REF][START_REF] Fredlund | The SpoIIQ landmark protein has different requirements for septal localization and immobilization[END_REF].

SpoIIQ (σ F -controlled) and the eight SpoIIIAA-AH proteins (σ E -controlled) form a complex known as the « feeding tube » that allows the MC to nurse the FS and to mantain a metabolic potential that still permits transcription and macromolecular synthesis after engulfment completion [START_REF] Camp | A feeding tube model for activation of a cell-specific transcription factor during sporulation in Bacillus subtilis[END_REF]. The feeding tube has homology to secretion systems and also transports small molecules into the FS that are necessary for σ G activation but remain unidentified [START_REF] Camp | A feeding tube model for activation of a cell-specific transcription factor during sporulation in Bacillus subtilis[END_REF][START_REF] Doan | Novel secretion apparatus maintains spore integrity and developmental gene expression in Bacillus subtilis[END_REF][START_REF] Fredlund | The SpoIIQ landmark protein has different requirements for septal localization and immobilization[END_REF]). Pre-engulfment activity of σ G in the FS is prevented by the FS anti-sigma factor, CsfB, which binds to and inactivates σ G [START_REF] Serrano | A negative feedback loop that limits the ectopic activation of a cell type-specific sporulation sigma factor of Bacillus subtilis[END_REF]. σ G transcribes 95 FS-specific genes encoding proteins that contribute to spore morphogenesis, germination and protection of DNA from damages (SASPs) [START_REF] Steil | Genome-wide analysis of temporally regulated and compartment-specific gene expression in sporulating cells of Bacillus subtilis[END_REF], Wang et al. 2006) [START_REF] Setlow | Absence of transient elevated UV resistance during germination of Bacillus subtilis spores lacking small, acid-soluble spore proteins alpha and beta[END_REF]). Among them, PdaA is required for modification of the cortex peptidoglycan [START_REF] Fukushima | A polysaccharide deacetylase gene (pdaA) is required for germination and for production of muramic deltalactam residues in the spore cortex of Bacillus subtilis[END_REF], the SpoVA is involved in dipicolinic acid (DPA) uptake and release, and GerA, GerB and GerK proteins are important for spore germination [START_REF] Paidhungat | Mechanisms of induction of germination of Bacillus subtilis spores by high pressure[END_REF]. σ G is also involved in the expression of genes coding for the membrane-embedded metalloprotease SpoIVB and CtpB involved in the processing and activation of σ K , the last sigma factor of sporulation (Figure 21) [START_REF] Campo | SpoIVB and CtpB are both forespore signals in the activation of the sporulation transcription factor sigmaK in Bacillus subtilis[END_REF]Rudner 2007, Mastny et al. 2013). Additionally, σ G transcribes the spoVT gene, coding for a transcriptional regulator that controls positively or negatively the expression of about half of the σ G -controlled genes (Figure 21) [START_REF] Bagyan | A compartmentalized regulator of developmental gene expression in Bacillus subtilis[END_REF], Wang et al. 2006). σ G together with SpoVT control the expression of ylyA gene, encoding the last FSspecific regulator. YlyA modulates the transcription of certain σ G -controlled genes, including genes involved in germination [START_REF] Traag | A novel RNA polymerase-binding protein controlling genes involved in spore germination in Bacillus subtilis[END_REF].

II.1.c. The mother cell line of gene expression: σ E and σ K

B. subtilis MC-specific σ factors are synthesized as inactive pro-proteins and activated by proteolytic cleavage [START_REF] Errington | Regulation of endospore formation in Bacillus subtilis[END_REF], Hilbert and Piggot 2004, Higgins and Dworkin 2012). σ E is the first σ factor to be activated in the MC compartment [START_REF] Labell | Sporulation-specific sigma factor sigma 29 of Bacillus subtilis is synthesized from a precursor protein, P31[END_REF]). Proσ E is encoded by spoIIGB, the second gene of the spoIIG operon whose transcription depends on a σ A -specific promoter that is activated by Spo0A-P [START_REF] Kenney | Gene encoding sigma E is transcribed from a sigma A-like promoter in Bacillus subtilis[END_REF]). Processing of pro-σ E into σ E in the MC is mediated by the membrane-bound SpoIIGA protease, encoded by the first gene of the spoIIG operon [START_REF] Stragier | Processing of a sporulation sigma factor in Bacillus subtilis: how morphological structure could control gene expression[END_REF]. This processing requires the product of the σ F -dependent spoIIR gene (Figure 21). The SpoIIR protein produced in the FS is localized in the intermembrane space, where it triggers the proteolysis of pro-σ E into σ E by SpoIIGA (Figure 21) [START_REF] Hofmeister | Extracellular signal protein triggering the proteolytic activation of a developmental transcription factor in B. subtilis[END_REF][START_REF] Karow | Identification of a gene, spoIIR, that links the activation of sigma E to the transcriptional activity of sigma F during sporulation in Bacillus subtilis[END_REF][START_REF] Fujita | An investigation into the compartmentalization of the sporulation transcription factor sigmaE in Bacillus subtilis[END_REF].

When activated, σ E controls the expression of 262 genes in the MC [START_REF] Eichenberger | The sigmaE regulon and the identification of additional sporulation genes in Bacillus subtilis[END_REF][START_REF] Steil | Genome-wide analysis of temporally regulated and compartment-specific gene expression in sporulating cells of Bacillus subtilis[END_REF]. The σ E -dependent genes are involved in the control of the FS engulfment through the transcription of three genes: spoIID, spoIIM and spoIIP [START_REF] Eichenberger | A three-protein inhibitor of polar septation during sporulation in Bacillus subtilis[END_REF]. SpoIID, SpoIIM and SpoIIP (DMP complex) form a PG degradation complex located at the engulfment membrane together with the PG synthesis machinery of the FS. in the FS through the synthesis of the SpoIIIAA-AH proteins (Aung et al. 2007, Chastanet and[START_REF] Chastanet | Engulfment during sporulation in Bacillus subtilis is governed by a multi-protein complex containing tandemly acting autolysins[END_REF] as well as both the synthesis and activation of the late MC sigma factor, σ K (Figure 21). σ E also transcribes the spoIIID and gerR genes encoding two transcriptional regulators (Figure 21) [START_REF] Kunkel | Temporal and spatial control of the mother-cell regulatory gene spoIIID of Bacillus subtilis[END_REF][START_REF] Eichenberger | The sigmaE regulon and the identification of additional sporulation genes in Bacillus subtilis[END_REF]. SpoIIID positively or negatively regulates σ E -controlled genes (Halberg andKroos 1994, Eichenberger et al. 2004),

while GerR acts as a repressor of some σ E -controlled genes and an activator of some σ Kdependent genes [START_REF] Eichenberger | The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis[END_REF][START_REF] Cangiano | Direct and indirect control of late sporulation genes by GerR of Bacillus subtilis[END_REF]. Moreover, SpoIIID together with σ E activates the transcription of the σ K encoding gene, sigK (Figure 21) [START_REF] Eichenberger | The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis[END_REF]). In fact, σ K is controlled both at the transcriptional and post-translational level. In B.

subtilis, as in other spore formers, the sigK gene is interrupted by a 48 kb prophage-like element called skin, which is excised during sporulation (Kunkel et al. 1990, Haraldsen andSonenshein 2003). Excision of the skin Bs is mediated by the site-specific recombinase, SpoIVCA, whose gene is present within the skin Bs element (Figure 22A) (Kunkel et al. 1990, Popham and[START_REF] Popham | Binding of the Bacillus subtilis spoIVCA product to the recombination sites of the element interrupting the sigma K-encoding gene[END_REF]. The spoIVCA gene, as sigK, is expressed under the dual control of σ E and SpoIIID in the MC limiting skin Bs excision to this compartment [START_REF] Eichenberger | The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis[END_REF]). After the skin excision, σ K is synthesized as an inactive percursor, pro-σ K , which will be then processed by SpoIVFB, a membrane-localized protease. SpoIVFB is itself helded inactive in a complex with two other membrane proteins SpoIVFA and BofA (Figure 21).

Removal of the pro-sequence requires a FS protein SpoIVB produced under σ G control (Figure 21). SpoIVB is localized into the intermembrane space and cleave the inhibitory protein SpoIVFA. SpoIVFB is thereby activated to process pro-σ K (Figure 21) [START_REF] Campo | SpoIVB and CtpB are both forespore signals in the activation of the sporulation transcription factor sigmaK in Bacillus subtilis[END_REF]. σ K control the expression of about 75 genes [START_REF] Eichenberger | The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis[END_REF][START_REF] Steil | Genome-wide analysis of temporally regulated and compartment-specific gene expression in sporulating cells of Bacillus subtilis[END_REF]. Most of them encode spore coat proteins or proteins required for spore maturation.

Among the members of the σ K regulon is the gene for the last regulator of the sporulation cascade, GerE. This regulator represses the transcription of over half of the σ K -target genes, while switching on 36 others (Figure 21) [START_REF] Zheng | Sporulation regulatory protein GerE from Bacillus subtilis binds to and can activate or repress transcription from promoters for mother-cell-specific genes[END_REF][START_REF] Eichenberger | The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis[END_REF].

II.2. Regulation of sporulation in clostridia

Even if the Spo0A regulator and the four cell-type sporulation specific σ factors σ F , σ E , σ G and σ K are present in clostridial genomes, substantial differences exist in the sporulation molecular programs between bacilli and clostridia as well as among clostridia (Paredes et al. 2005, Galperin et al. 2012, Abecasis et al. 2013, Saujet et al. 2013[START_REF] Al-Hinai | The Clostridium sporulation programs: diversity and preservation of endospore differentiation[END_REF], Fimlaid and Shen 2015). Genes encoding orthologues of the B. subtilis sensor HK, KinA-KinE, as well as the phosphorelay components Spo0B and Spo0F (Figure 20) are absent in clostridial genomes [START_REF] Worner | Phosphorylation and functional analysis of the sporulation initiation factor Spo0A from Clostridium botulinum[END_REF][START_REF] Steiner | Multiple orphan histidine kinases interact directly with Spo0A to control the initiation of endospore formation in Clostridium acetobutylicum[END_REF]. Furthermore, evidences point that phosphorylation of Spo0A results from two-component system activity rather than a multicomponent phosphorelay mechanism as observed in bacilli. Several orphan HKs that directly phosphorylate Spo0A have been identified in C. acetobutylicum [START_REF] Steiner | Multiple orphan histidine kinases interact directly with Spo0A to control the initiation of endospore formation in Clostridium acetobutylicum[END_REF],

C. difficile [START_REF] Underwood | Characterization of the sporulation initiation pathway of Clostridium difficile and its role in toxin production[END_REF] and C. botulinum [START_REF] Worner | Phosphorylation and functional analysis of the sporulation initiation factor Spo0A from Clostridium botulinum[END_REF] subtilis and a sigF mutant cannot initiate the asymmetric division [START_REF] Paredes | Transcriptional organization of the Clostridium acetobutylicum genome[END_REF], Jones et al. 2011). Moreover, σ K plays a dual role in sporulation, one before asymmetric division, and one in the very late stages of spore maturation [START_REF] Jones | The transcriptional program underlying the physiology of clostridial sporulation[END_REF][START_REF] Bi | SpoIIE is necessary for asymmetric division, sporulation, and expression of sigmaF, sigmaE, and sigmaG but does not control solvent production in Clostridium acetobutylicum ATCC 824[END_REF], Jones et al. 2011[START_REF] Al-Hinai | The Clostridium sporulation programs: diversity and preservation of endospore differentiation[END_REF]. Genome-wide analysis have demonstrated that in C. difficile, a weaker connection between the MC and the FS lines of gene expression exists when compared to B. subtilis (Pereira et al. 2013, Saujet et al. 2013).

II.2.a. Sporulation in C. difficile

Initiation of sporulation

In C. difficile, Spo0A plays a central role in the initiation of sporulation (Deakin et al. 2012) and controls the expression of 321 genes, involved not only in sporulation, but also in virulence and colonization [START_REF] Pettit | Functional genomics reveals that Clostridium difficile Spo0A coordinates sporulation, virulence and metabolism[END_REF]. spo0A is transcribed from both σ A -and σ Hdependent promoters (Saujet et al. 2011). As in B. subtilis, spo0A inactivation leads to an asporogenic phenotype [START_REF] Underwood | Characterization of the sporulation initiation pathway of Clostridium difficile and its role in toxin production[END_REF]. Five putative orphan HKs (CD1352, CD1492, CD1579, CD1949 and CD2492) are present in C. difficile genome. The HKs CD1492, CD1579 and CD2492 share some sequence identity with KinA-E of B. subtilis [START_REF] Underwood | Characterization of the sporulation initiation pathway of Clostridium difficile and its role in toxin production[END_REF]). CD1579 was shown to autophosphorylate and transfer its phosphate directly to Spo0A [START_REF] Underwood | Characterization of the sporulation initiation pathway of Clostridium difficile and its role in toxin production[END_REF]) and the inactivation of CD2492 decreases 3.5-fold sporulation efficiency [START_REF] Underwood | Characterization of the sporulation initiation pathway of Clostridium difficile and its role in toxin production[END_REF]). This suggests a role of both kinases in sporulation initiation (Figure 23). In contrast, CD1492 seems to have a negative effect on sporulation since its inactivation increases spore production [START_REF] Childress | The Phosphotransfer Protein CD1492 Represses Sporulation Initiation in Clostridium difficile[END_REF]). The signals or molecules triggering sporulation in C. difficile have not been identified yet.

However, environmental stimuli such as nutrient limitation and other stress factors might be involved. This is consistent with the impact of the CodY and CcpA regulators that negatively control sporulation in C. difficile (Figure 23) [START_REF] Dineen | Integration of metabolism and virulence by Clostridium difficile CodY[END_REF][START_REF] Antunes | Global transcriptional control by glucose and carbon regulator CcpA in Clostridium difficile[END_REF].

Indeed, both codY and ccpA mutants sporulate more effectively than the wild-type strain [START_REF] Antunes | Global transcriptional control by glucose and carbon regulator CcpA in Clostridium difficile[END_REF], Nawrocki et al. 2016). CcpA, the catabolite control protein, is implicated in the repression of genes involved in the control of the first stages of sporulation like spo0A and spoIIA operon [START_REF] Antunes | Global transcriptional control by glucose and carbon regulator CcpA in Clostridium difficile[END_REF], while CodY was found to negatively controls the expression of spo0A as well as the two HKs CD1579 and CD2492 (Figure 23) [START_REF] Dineen | Integration of metabolism and virulence by Clostridium difficile CodY[END_REF], Nawrocki et al. 2016). A study performed by Edwards et al suggested that C. difficile initiates sporulation in response to nutrient availability and uptake [START_REF] Edwards | Conserved oligopeptide permeases modulate sporulation initiation in Clostridium difficile[END_REF]. In agreement, an increased sporulation was observed when genes encoding two oligopeptide permeases, Opp and App, were inactivated, likely due to the inhability to import small peptides as a nutrient source. In other Gram positive bacteria, Opp and App are also involved in the import of small quorum sensing peptides, which control physiological processes such as sporulation, toxin expression and production of other virulence factors [START_REF] Perego | The oligopeptide transport system of Bacillus subtilis plays a role in the initiation of sporulation[END_REF][START_REF] Rudner | The spo0K locus of Bacillus subtilis is homologous to the oligopeptide permease locus and is required for sporulation and competence[END_REF][START_REF] Solomon | Purification and characterization of an extracellular peptide factor that affects two different developmental pathways in Bacillus subtilis[END_REF][START_REF] Gominet | Oligopeptide permease is required for expression of the Bacillus thuringiensis plcR regulon and for virulence[END_REF]). However, it remains unclear whether C. difficile employs quorum sensing to regulate sporulation initiation. Recently, the regulator RstA was found to positively control sporulation initiation in C. difficile, while repressing motility and toxin production (Figure 23) (Edwards et al. 2016b).

Sigma factors and transcriptional regulators

In B. subtilis, hierarchical regulatory cascades such as σ F ➝ RsfA/ σ G ➝ SpoVT➝ YlyA in the FS and σ E ➝ GerR/ SpoIIID ➝ σ K ➝ GerE in the MC (Figure 21) describe the programs of gene expression within each compartment [START_REF] Eichenberger | The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis[END_REF], Wang et al. 2006). These two lines of gene expression are connected by intercellular signalling pathways (Figure 21). In C. difficile, the sporulation σ factors exhibit compartment-specific activation similar to B. subtilis, with σ F and σ G activity being restricted to the FS and σ E and σ K activity being restricted to the MC (Fimlaid et al. 2013, Pereira et al. 2013). Similarly, the inactivation of sigF and sigE in C. difficile blocks the sporulation process after asymmetric division forming mutants with disporic forms. The sigG and sigK mutants are blocked after engulfment completion (Fimlaid et al. 2013, Pereira et al. 2013). sigG mutant does not make cortex while deposition of coat-like eletrodense material around the FS was observed contrary to what was detected in B. subtilis. C. difficile sigK mutant produces FS with a cortex layer but the coat layer is absent (Fimlaid et al. 2013, Pereira et al. 2013). This suggests that in C. (Fimlaid et al. 2013, Pereira et al. 2013, Saujet et al. 2013). Moreover, only two regulators, SpoVT in the FS and SpoIIID in the MC, are found in C. difficile (Pishdadian et al. 2015).

PDC

Compared to B. subtilis, in C. difficile (i) the σ E regulon is not strictly under the control of σ F although SpoIIR is essential for pro-

σ E cleavage; (ii) σ E is dispensable for σ G activation; (iii)
σ G is dispensable for σ K activation; and (iv) σ K does not undergo proteolytic activation in C.

difficile (Fimlaid et al. 2013, Pereira et al. 2013, Saujet et al. 2013).

The forespore regulatory network: σ F and σ G

The spoIIAA-spoIIAB-sigF operon is transcribed by σ H and positively controlled by Spo0A (Figure 24) (Saujet et al. 2011[START_REF] Rosenbusch | C. difficile 630Deltaerm Spo0A regulates sporulation, but does not contribute to toxin production, by direct high-affinity binding to target DNA[END_REF]. σ F , the first σ factor of the FS, is expressed before and after asymmetric division in both compartments, but the expression of its targets is observed only in the FS indicating that σ F is active only in the smaller compartment (Pereira et al. 2013). As in B. subtilis, the mechanism of activation of σ F is probably dependent on the anti-σ SpoIIAB, the anti-anti-σ factor SpoIIAA and the phosphatase SpoIIE (Figure 24) (Duncan andLosick 1993, Fimlaid andShen 2015). Once activated, σ F controls the expression of about 25 genes. Among them are spoIIR, spoIIQ and spoIIP. Similarly to B. subtilis, SpoIIQ and SpoIIP are critical for the engulfment completion (Serrano et al. 2016a[START_REF] Dembek | Peptidoglycan degradation machinery in Clostridium difficile forespore engulfment[END_REF][START_REF] Ribis | Differential requirements for conserved peptidoglycan remodeling enzymes during Clostridioides difficile spore formation[END_REF]). However, in B. subtilis, spoIIP is part of the σ E regulon. In C. difficile, σ F is also required to transcribe the gpr gene encoding a protease probably important for degradation of SASPs during germination [START_REF] Sussman | Cloning, nucleotide sequence, and regulation of the Bacillus subtilis gpr gene, which codes for the protease that initiates degradation of small, acid-soluble proteins during spore germination[END_REF], Fimlaid et al. 2013, Saujet et al. 2013). Moreover, σ F also controls the expression of genes thought to be involved in PG metabolism: sleB, CD2141 and CD1229 encoding a spore cortex hydrolysing enzyme, a DAla-DAla carboxypeptidase and a PG glycosyltransferase, respectively. Like in B. subtilis, the spoIIGA, sigE and sigG genes are clustered in C. difficile (Figure 24) [START_REF] De Hoon | Hierarchical evolution of the bacterial sporulation network[END_REF]. In C. difficile, sigG gene is likely transcribed from two promoters, one recognized by σ A upstream of the spoIIGA gene and one upstream of the sigG gene recognized by σ F and/or σ G , since a promoter with both FS σ factor consensus sequences have been mapped (Saujet et al. 2011, Saujet et al. 2013). In B. subtilis, the onset of σ G activity coincides with engulfment completion and requires the activity of σ E .

However, in C. difficile, σ G activation does not require engulfment to be completed and is independent of σ E (Pereira et al. 2013, Saujet et al. 2013) suggesting that the feeding tube is dispensable for σ G activity. In C. difficile, since a sigF mutant produces wild-type levels of σ G but decreased levels of the σ G -target genes, σ G seems to require σ F for post-translational activation via an unknown mechanism (Figure 24) (Fimlaid et al. 2013). σ G controls the expression of 50 FS-specific genes including genes encoding the SASPs and the spoVA operon, which is required for the import of DPA from the MC to the FS (Donnelly et al. 2016). Additionally, genes encoding proteins that might be involved in protection against oxidative stress are controlled by σ G . This includes CD1567 (CotG), CD1631 (SodA) and CD2845, which encode a catalase, a SOD and a rubrerythrin, respectively, as mentioned earlier in Chapter 3 (Permpoonpattana et al. 2013, Saujet et al. 2013). Genes involved in cell wall synthesis and cortex formation are also controlled by this σ factor (Fimlaid et al. 2013, Saujet et al. 2013). CD1430 (PdaA1) encodes a N-acetylmuramic acid deacetylase involved in the formation of muramic-δ-lactam in the spore cortex PG [START_REF] Coullon | N-Deacetylases required for muramic-delta-lactam production are involved in Clostridium difficile sporulation, germination and heat resistance[END_REF][START_REF] Diaz | Clostridium difficile Lipoprotein GerS Is Required for Cortex Modification and Thus Spore Germination[END_REF]). In addition, CD1291 (dacF), wich encodes a D-Ala-D-Ala carboxypeptidase likely regulating the degree of cross-linking of the spore PG was also positively controlled by σ G .

Finally, the spoVT gene encoding the SpoVT regulator is probably transcribed under the dual control of σ F and σ G (Fimlaid et al. 2013, Saujet et al. 2013). SpoVT is involved in the regulation of σ G -targets such as sspA and sspB encoding the SASPs (Figure 24). Additionally, SpoVT seems to negatively regulate spoIIR and gpr, suggesting that, in C. difficile, this regulator might play the role of RsfA in the negative control of σ F targets (Saujet et al. 2013).

The mother cell regulatory network: σ E and σ K

The sigE gene is transcribed from a σ A -dependent promoter and positively controlled by Spo0A (Figure 24) (Saujet et al. 2011[START_REF] Rosenbusch | C. difficile 630Deltaerm Spo0A regulates sporulation, but does not contribute to toxin production, by direct high-affinity binding to target DNA[END_REF]. In C. difficile, σ F was found to positively regulate the expression of spoIIR. Whereas in B. subtilis σ E maturation in the MC after asymmetric division is strictly dependent on σ F , in C. difficile, such maturation is partially independent of this FS σ factor. Indeed, in C. difficile the synthesis of SpoIIR, that triggers the proteolysis of pro-σ E into σ E by SpoIIGA is partially independent on σ F (Figure 24) (Saujet et al. 2013). The σ E regulon comprises around 97 genes and is the largest regulon among the four sporulation specific σ factors as observed in B. subtilis. σ E transcribes genes encoding proteins involved in engulfment control, formation of the feeding tube, synthesis of the cortex, germination and initiation of coat assembly (Fimlaid et al. 2013, Saujet et al. 2013). The spoIID gene encoding a PG hydrolase required for the engulfment of the FS is controlled by σ E as well as the cwlD gene, encoding a N-acetylmuramoyl-L-alanine amidase that together with the FS-controlled PdaA1 generates muramic-δ-lactam (Saujet et al. 2013[START_REF] Diaz | Clostridium difficile Lipoprotein GerS Is Required for Cortex Modification and Thus Spore Germination[END_REF]. As in B. subtilis, σ E activates the expression of the spoIIIAA operon. The genes of the csp operon, encoding the germinant receptor (CspC) and the proteases responsible for the activation of cortex-lytic enzymes (CspAB) during germination, are also under the control of σ E (Adams et al. 2013, Francis et al. 2013a, Saujet et al. 2013).

Addtionally, a σ E -dependent promoter was mapped upstream of spoIVA, sipL and cotB genes, all encoding spore coat proteins (Permpoonpattana et al. 2011a, Putnam et al. 2013, Saujet et al. 2013). Finally, σ E transcribes the genes encoding σ K and the transcriptional regulator SpoIIID, which negatively controls the transcription of members of the σ E regulon (such as spoIVA, sipL, cotB and the spoIIIA operon) and positively controls the expression of some members of the σ K regulon (Figure 24) (Fimlaid et al. 2013, Pereira et al. 2013). In C.

difficile, the sigK gene is transcribed from two different promoters: one dependent on σ E and the other dependent on σ K and positively controlled by SpoIIID (Fimlaid et al. 2013, Pereira et al. 2013, Saujet et al. 2013, Pishdadian et al. 2015). The positive control of SpoIIID under σ K -controlled genes is due either to a direct binding of the regulator or via the positive control that SpoIIID exerces on sigK transcription. While σ K is synthesized in B. subtilis as an inactive pre-protein, σ K of C. difficile lacks a pro-sequence and the cleavage proteins (Figure 24 and 22B). Therefore, in C. difficile, σ K is produced directly in its mature form.

Accordingly, the absence of a cleavage process at the end of the engulfment, avoids a dependency of σ G on σ K activity in this bacterium (Pereira et al. 2013, Saujet et al. 2013).

The skin Cd element

As observed in B. subtilis, a skin Cd element is also present in the sigK gene of C. difficile (Figure 22B) [START_REF] Stragier | Chromosomal rearrangement generating a composite gene for a developmental transcription factor[END_REF][START_REF] Kunkel | The Bacillus subtilis gene for the development transcription factor sigma K is generated by excision of a dispensable DNA element containing a sporulation recombinase gene[END_REF][START_REF] Haraldsen | Efficient sporulation in Clostridium difficile requires disruption of the sigmaK gene[END_REF]. Compared with B. subtilis, the skin Cd differs in size, gene content, site of insertion and orientation suggesting independent integrations into sigK gene during evolution [START_REF] Haraldsen | Efficient sporulation in Clostridium difficile requires disruption of the sigmaK gene[END_REF]. The skin Cd element has 19 open reading frames and a cassette of the CRISPR system used for protection against foreign DNA [START_REF] Monot | Reannotation of the genome sequence of Clostridium difficile strain 630[END_REF], Soutourina et al. 2013). Two proteins, CD1231 and CD1234, encoded by genes located within the skin Cd are required for the excision of this element (Figure 22B) (Serrano et al. 2016b

SpoVA

dependent on both σ E and SpoIIID which results in the production of CD1234 only in the MC (Serrano et al. 2016b). The late production of CD1234 delays and restricts skin Cd excision to the terminal MC. Thus, in C. difficile, the crucial element regulating σ K activity is the skin Cd excision (Serrano et al. 2016b). Then, the sigK gene is reconstituted, transcribed and translated. Global transcription profiling revealed that σ K controls the transcription of 56 genes encoding proteins involved in processes such as synthesis of DPA, the assembly of the spore coat and exosporium as well as in the lysis of the MC (Figure 24) (Fimlaid et al. 2013, Saujet et al. 2013). The dpaA and dpaB genes encoding the DPA synthase are controlled by σ K (Donnelly et al. 2016). In addition, genes encoding coat proteins such as CotA, CotE or the YabG protease are also positively controlled by σ K . This is also the case for the bclA1, bclA2

and bclA3 genes encoding exosporium proteins and of two genes encoding N-acetylmuramic acid L-alanine amidases (CD1898 and CD2184) that are possibly involved in MC lysis.

Finally, sleC encoding the major spore cortex lytic enzyme is also a member of σ K regulon.

III. Germination

The process by which the spores reinitiate growth is called germination. Germination usually occurs in response to environmental signals including amino acids and cell wall PG muropeptides derived from growing cells (Setlow 2003[START_REF] Shah | A eukaryotic-like Ser/Thr kinase signals bacteria to exit dormancy in response to peptidoglycan fragments[END_REF]). Since CDI is mediated by toxin producing vegetative cells, germination of C. difficile spores is an important initial step for the disease process [START_REF] Howerton | A new strategy for the prevention of Clostridium difficile infection[END_REF]. Indeed, C. difficile strains presenting a defected germination are associated to a decreased virulence (Francis et al. 2013a, Fimlaid et al. 2015).

III.1. Germination in B. subtilis

In B. subtilis, nutrient germinants are sensed by Ger-type germinant receptors (GRs) synthesized in the FS under σ G control, and localized at the inner membrane of the spore (Figure 25) (Paredes-Sabja et al. 2011). The germinants must pass through the spore coat, the FS outer membrane and the cortex to access the GRs. Thus, it is possible that there are specialized mechanisms to facilitate germinants access into the spore. In some bacilli, the GerP proteins located in the spore coat may play this function [START_REF] Carr | Role of the gerP operon in germination and outgrowth of Bacillus anthracis spores[END_REF][START_REF] Butzin | Analysis of the effects of a gerP mutation on the germination of spores of Bacillus subtilis[END_REF]. In bacilli and some clostridia, the binding of the germinant to the GRs, triggers the release of the DPA chelated to calcium (Ca-DPA) and of the monovalent ions (H + , Na + and K + ) stored in the spore core. During sporulation, DPA is initially synthesized in the MC (Daniel andErrington 1993, Orsburn et al. 2010) and then transported into the core of the FS by a two step transport mechanism. DPA is first translocated into the intermembrane space by the FS outer membrane protein SpoVV (ylbJ, a σ E regulated gene) and then is transfered into the spore core by the SpoVA complex located in the inner spore membrane [START_REF] Tovar-Rojo | The products of the spoVA operon are involved in dipicolinic acid uptake into developing spores of Bacillus subtilis[END_REF][START_REF] Ramirez-Guadiana | A two-step transport pathway allows the mother cell to nurture the developing spore in Bacillus subtilis[END_REF]. The same SpoVA channel is involved in the release of Ca-DPA during germination [START_REF] Vepachedu | Role of SpoVA proteins in release of dipicolinic acid during germination of Bacillus subtilis spores triggered by dodecylamine or lysozyme[END_REF]. In B. subtilis, the Ca-DPA released binds to the cortex hydrolase CwlJ as a cofactor leading to the CwlJ activity (Figure 25) [START_REF] Paidhungat | Genetic requirements for induction of germination of spores of Bacillus subtilis by Ca(2+)-dipicolinate[END_REF]. SleB is other cortex lytic enzyme which seems to be activated by cortex deformation [START_REF] Paidhungat | Genetic requirements for induction of germination of spores of Bacillus subtilis by Ca(2+)-dipicolinate[END_REF]. The cortex degradation allows core hydration and resumption of metabolism in this compartment that results in the formation of the vegetative cell (Paidhungat et al. 2001, Vepachedu and[START_REF] Setlow | I will survive: DNA protection in bacterial spores[END_REF].

III.2. Germination in C. difficile

C. difficile genome lacks orthologs of GerA, GerB and GerK receptors involved in germination in B. subtilis [START_REF] Sebaihia | The multidrug-resistant human pathogen Clostridium difficile has a highly mobile, mosaic genome[END_REF] suggesting that this bacterium responds to different environmental signals that those identified in bacilli. In fact, C. difficile spores germinate in response to a combination of specific bile salts (i.e., cholate and its derivatives [taurocholate, glycocholate and deoxycholate]). In addition, both L-glycine and divalent cations such as Ca 2+ and Mg 2+ can also act as co-germinants [START_REF] Kochan | Intestinal calcium and bile salts facilitate germination of Clostridium difficile spores[END_REF]. A recent study showed that Ca 2+ present within the gastrointestinal tract play a major role in inducing germination in vivo. Ca 2+ depletion of the ileal mouse contents reduced spore germination from 630 C. difficile strain by 90% [START_REF] Kochan | Intestinal calcium and bile salts facilitate germination of Clostridium difficile spores[END_REF]). The C. difficile genome encodes an operon of subtilisin-like proteases, cspBAC, controlled by σ E that is essential for germination (Saujet et al. 2013, Kevorkian andShen 2017). CspC was identified as the bile salt GR of C.

difficile and it is located in the spore coat/ FS outer membrane (Figure 26) (Francis et al.

(Figure 26). CspB is required for the activation of the cortex lytic enzyme SleC, by cleavage of the inactive pro-SleC into SleC. Once activated, SleC initiates degradation of the cortex PG (Figure 26) [START_REF] Burns | SleC is essential for germination of Clostridium difficile spores in nutrient-rich medium supplemented with the bile salt taurocholate[END_REF], Adams et al. 2013[START_REF] Kevorkian | Revisiting the Role of Csp Family Proteins in Regulating Clostridium difficile Spore Germination[END_REF]. SleC is thought to recognize the cortex-specific modification muramic-δ-lactam [START_REF] Burns | SleC is essential for germination of Clostridium difficile spores in nutrient-rich medium supplemented with the bile salt taurocholate[END_REF][START_REF] Gutelius | Functional analysis of SleC from Clostridium difficile: an essential lytic transglycosylase involved in spore germination[END_REF]. Contrary to B. subtilis, the hydrolysis of the cortex in C. difficile, precedes DPA release (Francis et al. 2015, Wang et al. 2015). Swelling of the core after cortex hydrolysis leads to DPA release through SpoVAC most likely by a mechanosensing mechanism [START_REF] Francis | Dipicolinic Acid Release by Germinating Clostridium difficile Spores Occurs through a Mechanosensing Mechanism[END_REF]. The role of the co-germinants and the events that lead to CspB activation are the major weakness of the model. A study showed that Ca 2+ is required for CspB-mediated SleC cleavage [START_REF] Kochan | Intestinal calcium and bile salts facilitate germination of Clostridium difficile spores[END_REF]. Ca 2+ from the spore core or from the external environment might bind to and activates CspB, with subsequent SleC activation and cortex hydrolysis. Another hypothesis is that CspC activates CspB by protein-protein interaction.

Other proteins like GerG, GerS and CD3298 are also involved in the germination process. All of them are encoded by genes controlled by σ E . GerS is a lipoprotein recently shown to be necessary for CwlD-mediated cortex modification (Fimlaid et al. 2015[START_REF] Diaz | Clostridium difficile Lipoprotein GerS Is Required for Cortex Modification and Thus Spore Germination[END_REF].

GerS and CwlD directly interact and CwlD mediates GerS incorporation into mature spores.

The cortex modifications are required for an efficient cortex hydrolysis by the lytic enzyme SleC [START_REF] Diaz | Clostridium difficile Lipoprotein GerS Is Required for Cortex Modification and Thus Spore Germination[END_REF]. Another protein required for C. difficile germination is GerG. Since gerG mutant produces spores lacking Csp proteins, the hypothesis is that GerG is required for optimal incorporation of CspA, CspB and CspC into spores [START_REF] Donnelly | A Clostridium difficile-Specific, Gel-Forming Protein Required for Optimal Spore Germination[END_REF]. CD3298 is an ATPase likely involved in transport of Ca-DPA across the FS outer membrane [START_REF] Kochan | Intestinal calcium and bile salts facilitate germination of Clostridium difficile spores[END_REF]. The overall architecture of the spore is conserved among endospore-forming bacteria.

A C

The spore is a spherical or roughly ovoid structure composed by concentric layers, surrounding the core in its center (Figure 27A). The spore core harbors the chromosomic DNA, maintained in a highly compact state by the SASPs that are believed to block transcription and protect the DNA [START_REF] Setlow | I will survive: DNA protection in bacterial spores[END_REF][START_REF] Moeller | Roles of the major, small, acid-soluble spore proteins and sporespecific and universal DNA repair mechanisms in resistance of Bacillus subtilis spores to ionizing radiation from X rays and high-energy charged-particle bombardment[END_REF][START_REF] Paredes-Sabja | Role of small, acid-soluble spore proteins in the resistance of Clostridium perfringens spores to chemicals[END_REF][START_REF] Setlow | Spore Resistance Properties[END_REF]. The core also contains DPA in a 1:1 complex with divalent cations, mostly Ca 2+ . The core is delimited by an inner membrane that retards the diffusion of water and serves as permeability barrier against chemicals (Setlow 2003[START_REF] Sunde | The physical state of water in bacterial spores[END_REF][START_REF] Knudsen | Water and Small-Molecule Permeation of Dormant Bacillus subtilis Spores[END_REF]. In many species, this membrane houses the germinant receptors (Paredes-Sabja et al. 2011). The inner membrane will become the plasma membrane after germination. Around the FS inner membrane is a PG-containing germ cell wall that will become the cell wall for the emerging cell during outgrowth. The germ cell wall is surrounded by the spore cortex layer and the outer membrane. The cortex plays a major role in mantaining the dehydrated state of the core by constricting the core's volume. The cortex is a modified PG-containing layer, where the N-acetyl muramic acid is replaced by muramic-δlactam [START_REF] Meador-Parton | Structural analysis of Bacillus subtilis spore peptidoglycan during sporulation[END_REF]. This modification is essential for specific hydrolysis by cortex lytic enzymes during germination, which degrade only the cortex layer and not the germ cell wall. Concomitantly with cortex formation, the proteinaceous spore coat layer is deposited at the surface of the outer membrane. This layer is universally present but varies in composition and morphology among species [START_REF] Aronson | Structure and morphogenesis of the bacterial spore coat[END_REF], Henriques and Moran 2000[START_REF] Giorno | Morphogenesis of the Bacillus anthracis spore[END_REF][START_REF] Driks | The Spore Coat[END_REF]). In the case of B. subtilis, the spore coat is enfolded by the crust (Figure 27B) [START_REF] Mckenney | A distance-weighted interaction map reveals a previously uncharacterized layer of the Bacillus subtilis spore coat[END_REF]. In other species such as B. anthracis and C. difficile, the most external spore layer is the exosporium (Figure 27C) [START_REF] Stewart | The Exosporium Layer of Bacterial Spores: a Connection to the Environment and the Infected Host[END_REF]. Most of these spore structures are rapidly lost during germination i.e., the cortex is degraded by cell wall hydrolases and the coat is shed.

The fate of the outer spore membrane is less clear.

II. Spore resistance

Spores are metabolically dormant and partially dehydrated, which are likely characteristics that allow their survival in nutrient-free and harsh environments [START_REF] Setlow | Spores of Bacillus subtilis: their resistance to and killing by radiation, heat and chemicals[END_REF][START_REF] Setlow | Spore Resistance Properties[END_REF]. They exhibit extraordinary resistance properties including resistance to ultraviolet (UV) radiation, chemicals (such as H 2 O 2 and hypochlorite), extreme heat and other stresses.

Therefore, high level of Ca-DPA and partial dehydration of the spore core, a functional cortex, RecA-dependent DNA repair machinery and SASPs are important requirements for resistance to extreme heat [START_REF] Setlow | Spores of Bacillus subtilis: their resistance to and killing by radiation, heat and chemicals[END_REF][START_REF] Setlow | I will survive: DNA protection in bacterial spores[END_REF][START_REF] Paredes-Sabja | Role of small, acid-soluble spore proteins in the resistance of Clostridium perfringens spores to chemicals[END_REF], Donnelly et al. 2016). During late stages of sporulation, most of the water in the spore core is replaced with Ca-DPA, and mutants deficient in Ca-DPA do not survive desiccation.

II.1. Functions of the spore coat

The coat protects against a wide range of assaults, including digestion by predatory microbes and challenges by various chemicals [START_REF] Nicholson | Resistance of Bacillus endospores to extreme terrestrial and extraterrestrial environments[END_REF][START_REF] Riesenman | Role of the spore coat layers in Bacillus subtilis spore resistance to hydrogen peroxide, artificial UV-C, UV-B, and solar UV radiation[END_REF][START_REF] Klobutcher | The Bacillus subtilis spore coat provides "eat resistance" during phagocytic predation by the protozoan Tetrahymena thermophila[END_REF][START_REF] Setlow | Spore Resistance Properties[END_REF]. For example, intact B. subtilis spores can survive ingestion by Tetrahymena thermophila. However, coat disruption results in a digestion of the cortex by the protozoa [START_REF] Klobutcher | The Bacillus subtilis spore coat provides "eat resistance" during phagocytic predation by the protozoan Tetrahymena thermophila[END_REF]). Resistance to these organisms is likely due to exclusion of microbial degradative enzymes by sieving. For instance, the spore coat protects the underlying spore cortex, which is susceptible to PG-degrading enzymes such as lysozyme.

It seems reasonable to assume that the coat acts as a barrier, passively excluding degradative enzymes [START_REF] Setlow | Spores of Bacillus subtilis: their resistance to and killing by radiation, heat and chemicals[END_REF]). However, the spore coat is not impermeable, because germinants can pass through it en route to the GRs in the inner membrane [START_REF] Setlow | Germination of Spores of the Orders Bacillales and Clostridiales[END_REF]. The coat also protects against several types of reactive chemicals, such as chlorine dioxide, hypochlorite, ozone, peroxynitrite and H 2 O 2 [START_REF] Setlow | Spores of Bacillus subtilis: their resistance to and killing by radiation, heat and chemicals[END_REF][START_REF] Setlow | Spore Resistance Properties[END_REF]. B. subtilis spores lacking the CotE protein essential for coat morphogenesis are more sensitive to H 2 O 2 and hypochlorite [START_REF] Riesenman | Role of the spore coat layers in Bacillus subtilis spore resistance to hydrogen peroxide, artificial UV-C, UV-B, and solar UV radiation[END_REF]Nicholson 2000, Henriques andMoran 2007). Therefore, more specific resistance properties, like those directed against oxidative compounds, might be provided by catalases and other enzymes present in the coat. A Mn 2+ -dependent SOD (SodA) and at least two Mn 2+ pseudocatalases, CotJC and YjqC, are associated with the B. subtilis coat layers. However, a possible role in protection against ROS has never been shown [START_REF] Henriques | Characterization of cotJ, a sigma E-controlled operon affecting the polypeptide composition of the coat of Bacillus subtilis spores[END_REF][START_REF] Henriques | Involvement of superoxide dismutase in spore coat assembly in Bacillus subtilis[END_REF]. In C. difficile, CotCB, CotD and CotG catalases are present in the surface layers of spores and cotD mutant spores showed a reduction in catalase activity compared to the wild-type strain (Permpoonpattana et al. 2013).

Protection against UV radiation and H 2 O 2 seems to be provided in part by CotA, a copper-dependent laccase present in the outer coat of B. subtilis, which generates a melanin-like dark brown pigment that accumulates in the spores [START_REF] Hullo | CotA of Bacillus subtilis is a copper-dependent laccase[END_REF][START_REF] Martins | Molecular and biochemical characterization of a highly stable bacterial laccase that occurs as a structural component of the Bacillus subtilis endospore coat[END_REF]. In addition, spore coat components might react with and detoxify some chemicals before they reach and attack their targets further within the spore [START_REF] Riesenman | Role of the spore coat layers in Bacillus subtilis spore resistance to hydrogen peroxide, artificial UV-C, UV-B, and solar UV radiation[END_REF], Henriques and Moran 2007[START_REF] Setlow | Spore Resistance Properties[END_REF]).

An integral coat seems to be also required for an efficient germination. In B. subtilis, the assembly of the cortex lytic enzyme CwlJ is dependent on the coat protein GerQ and the spores of the gerQ mutant germinate poorly [START_REF] Bagyan | Localization of the cortex lytic enzyme CwlJ in spores of Bacillus subtilis[END_REF]. The inactivation of the gerP operon, mentioned earlier in the Chapter 4, section III.1 Germination in B. subtilis, also blocks germination prior to cortex hydrolysis in B. subtilis and B. cereus. GerP proteins seems to play a crucial role producing a permeable coat to small hydrophilic germinants to gain access to their receptors in the inner layers of the spore [START_REF] Behravan | Mutations in the gerP locus of Bacillus subtilis and Bacillus cereus affect access of germinants to their targets in spores[END_REF][START_REF] Butzin | Analysis of the effects of a gerP mutation on the germination of spores of Bacillus subtilis[END_REF].

III. Spore surface layers of bacilli and clostridia

III.1. Conservation of proteins

The most internal spore compartments, core and cortex, do not usually display interspecies differences that are distinguishable by electron microscopy. Moreover, genes encoding proteins involved in the formation of the spore cortex, spoVB, spoVD, spoVE, yabP, yabQ, ylbJ, yqfC and yqfD in B. subtilis are conserved in spore forming bacteria, suggesting a conservation of the cortex biosynthesis [START_REF] Fawcett | The transcriptional profile of early to middle sporulation in Bacillus subtilis[END_REF][START_REF] Asai | The Bacillus subtilis yabQ gene is essential for formation of the spore cortex[END_REF][START_REF] Eichenberger | The sigmaE regulon and the identification of additional sporulation genes in Bacillus subtilis[END_REF]. By contrast, considerable diversity is observed in the structure of the most external layers such as the spore coat. About 80% of the spore coat proteins are conserved between B. subtilis and B. anthracis. In addition, 16 coat proteins appear to be unique to B. subtilis such as the cgeAB and cgeCDE operons encoding spore coat maturation proteins (Henriques and Moran 2007). In clostridia, conservation of B. subtilis coat proteins is even more limited (Henriques and Moran 2007[START_REF] De Hoon | Hierarchical evolution of the bacterial sporulation network[END_REF], Permpoonpattana et al. 2011b, Galperin et al. 2012). Only about 20 genes of the B. subtilis encoding coat structural proteins have their homologs in the sequenced clostridial genomes. Considering that the spore surface represents the interface between spores and the environment, and given the diversity in coat structure and composition among species, it seems reasonable to speculate that this diversity is driven by adaptation of spore forming bacteria to variety of niches.

III.2. Spore coat organization in B. subtilis

The spore coat of B. subtilis is a multilayered proteinaceous structure, composed at least by 70 proteins with a range in size from 6 kDa to larger than 70 kDa (Henriques and Moran 2007, McKenney et al. 2013[START_REF] Driks | The Spore Coat[END_REF]. Proteins found in the coat layers can be extracted from purified spores by treatment with reducing agents in the presence of detergents (Henriques and Moran 2007). This spore layer is divided in an undercoat or basement layer, a lamellar inner coat and an electrodense outer coat (Figure 28) (McKenney and Eichenberger 2012). The inner and outer coat gained these designations due to the differences observed using TEM and osmium tetroxide staining i.e., the inner coat was more lightly and contained multiple concentric dark lamellae and the outer coat was electrodense and darkly stained [START_REF] Warth | The composition and structure of bacterial spores[END_REF][START_REF] Giorno | Morphogenesis of the Bacillus anthracis spore[END_REF], Henriques and Moran 2007). Both coat layers are packed closely together and appear thicker at the spore poles. The factors that determine the pattern of each layer is not known but most likely reflects the successive overlay of the coat proteins during the assembly and their physical properties. The basement layer, is not distinguishable by TEM, but corresponds to the innermost layer atop which the rest of the coat assembles. It contains several coat proteins required to initiate coat assembly such as SpoIVA, SpoVM and SpoVID (Figure 28). These proteins help bridging the coat to the cortex layer [START_REF] Driks | The Spore Coat[END_REF]. Indeed, some mutants with deficiencies in coat assembly (e.g., cotE) have undercoat regions that do not properly adhere to the underlying cortex (Zheng et al. 1988[START_REF] Aronson | Protein filaments may initiate the assembly of the Bacillus subtilis spore coat[END_REF][START_REF] Driks | The Spore Coat[END_REF].

III.2.a. Coat morphogenetic proteins

In B. subtilis, the formation of the coat layers is governed by the morphogenetic factors SpoVM, SpoIVA, SpoVID, SafA, and CotE (Figure 28) (Henriques andMoran 2007, McKenney et al. 2013). They were named morphogenetic proteins as they play a major role in coat morphogenesis without affecting gene expression. A combination of genetic and electron microscopy analyses was used to infere the locations of the morphogenetic proteins. Only

SpoIVA is present in all bacilli and clostridia. SpoIVA is produced in the MC under the control of σ E , and is essential for the assembly of both the spore cortex and coat layers [START_REF] Piggot | Genetic aspects of bacterial endospore formation[END_REF], Roels et al. 1992[START_REF] Stevens | Characterization of a sporulation gene, spoIVA, involved in spore coat morphogenesis in Bacillus subtilis[END_REF][START_REF] Catalano | Amino acids in the Bacillus subtilis morphogenetic protein SpoIVA with roles in spore coat and cortex formation[END_REF]. SpoIVA binds and hydrolyses ATP to form highly stable polymers that encase the FS (Ramamurthi andLosick 2008, Wu et al. 2015). The localization of SpoIVA requires SpoVM, also produced under the control of σ E (Figure 28) [START_REF] Price | A four-dimensional view of assembly of a morphogenetic protein during sporulation in Bacillus subtilis[END_REF]. Both proteins were find to localize early to the MC face of the FS outer membrane and were observed tracking along with the engulfing FS membrane forming a full spherical shell that encases the spore in a step known as "spore encasement" [START_REF] Driks | Subcellular localization of proteins involved in the assembly of the spore coat of Bacillus subtilis[END_REF][START_REF] Pogliano | Visualization of the subcellular location of sporulation proteins in Bacillus subtilis using immunofluorescence microscopy[END_REF][START_REF] Price | A four-dimensional view of assembly of a morphogenetic protein during sporulation in Bacillus subtilis[END_REF], van Ooij and Losick 2003). To this end, the C-terminal part of SpoIVA interacts with the Nterminal region of SpoVM and then polymerizes and surrounds the FS (Figure 28) [START_REF] Ramamurthi | Peptide anchoring spore coat assembly to the outer forespore membrane in Bacillus subtilis[END_REF]. SpoVM is a 26-amino-acid-long protein that recognizes the positive curvature of the FS outer membrane and embeds itself there [START_REF] Levin | An unusually small gene required for sporulation by Bacillus subtilis[END_REF][START_REF] Ramamurthi | Geometric cue for protein localization in a bacterium[END_REF][START_REF] Gill | Structural basis for the geometry-driven localization of a small protein[END_REF]. Both spoIVA and spoVM mutants fail to make cortex, and the coat accumulates as swirls floating in the MC cytoplasm [START_REF] Roels | Characterization of spoIVA, a sporulation gene involved in coat morphogenesis in Bacillus subtilis[END_REF][START_REF] Levin | An unusually small gene required for sporulation by Bacillus subtilis[END_REF]. Even if coat assembly and cortex formation are events that occur in different compartments, MC and intermembrane space respectively, SpoIVA and SpoVM are required for both processes. At this initial stage of coat formation, a quality control mechanism is present that prevents cortex formation if SpoIVA fails to encase the FS (Tan et al. 2015). It is dependent on the CmpA protein and the ClpXP protease. In the presence of spore coat defects, CmpA binds to improperly localized SpoIVA and delivers it to ClpXP for degradation [START_REF] Tan | A Quality-Control Mechanism Removes Unfit Cells from a Population of Sporulating Bacteria[END_REF]. After elimination of SpoIVA, cortex formation is inhibited and the sporulating cell eventually lyses. In the absence of defect, CmpA is eliminated by ClpXPmediated proteolysis. SpoIVA and SpoVM finish spore encasement as soon as engulfment is completed (McKenney and Eichenberger 2012). However, SpoIVA does not complete encasement in the absence of another coat morphogenetic protein SpoIVD (Wang et al. 2009a) whose production is also under the control of σ E (Beall et al. 1993). SpoIVA proper assembly requires SpoIVD, but SpoIVA is also required for the localization of SpoVID to the surface of the developing spore (Figure 28) (Beall et al. 1993[START_REF] Driks | Subcellular localization of proteins involved in the assembly of the spore coat of Bacillus subtilis[END_REF]. This mutual dependency is supported by the evidence that SpoIVA interacts with the SpoVID's N terminus (Figure 28). The phenotype of spoVID mutant cells is reminescent of the observed in the spoIVA and spoVM mutants (Beall et al. 1993). However, SpoVID seems to be specifically devoted to the spore coat layer, since spoVID mutant is able to form the cortex layer (Beall et al. 1993). The swirls of coat material that accumulate in the MC cytoplasm of these three mutants, suggests that coat proteins are capable of self assembly even in the absence of the SpoIVA, SpoVM and SpoIVD localization [START_REF] Aronson | Protein filaments may initiate the assembly of the Bacillus subtilis spore coat[END_REF]. Therefore, the assembly of the basement layer of the coat requires the combined contributions of these three proteins (Figure 28). Assembly of the coat layers beyond the basement layer is dependent on additional morphogenetic proteins [START_REF] Driks | Subcellular localization of proteins involved in the assembly of the spore coat of Bacillus subtilis[END_REF]). Among them, SafA and CotE are respectively the morphogenetic proteins of the inner and outer coat (Figure 28) and are produced under the control of σ E , while cotE has also a promoter recognized by σ K (Zheng et al. 1988[START_REF] Ozin | Morphogenetic proteins SpoVID and SafA form a complex during assembly of the Bacillus subtilis spore coat[END_REF]. Recruitment of both SafA and CotE is dependent on SpoIVA and SpoVID (Figure 28) [START_REF] Mullerova | Interactions between Bacillus subtilis early spore coat morphogenetic proteins[END_REF], de Francesco et al. 2012). An interaction between SafA and CotE has not been observed. SafA, located in the inner coat (Figure 28) (Takamatsu et al. 1999) interacts with both SpoIVA and SpoVID but the interaction with this last protein is stronger [START_REF] Ozin | Morphogenetic proteins SpoVID and SafA form a complex during assembly of the Bacillus subtilis spore coat[END_REF], Mullerova et al. 2009). In the spore, CotE is located at the interface between the inner and outer coats (Figure 28) [START_REF] Driks | Subcellular localization of proteins involved in the assembly of the spore coat of Bacillus subtilis[END_REF]). The assembly of the two coat layers seems to occur independently, since safA mutant spores lack an inner coat but retain a seemingly unaffected outer coat and cotE mutant spores retain the lamellar structure of the inner coat but lack an outer coat (Zheng et al. 1988, Takamatsu et al. 1999, Henriques and Moran 2007). However, CotE appears to contribute to the attachment of the coat to the spore surface. The assembly of the outer coat involves the cooperation of CotE with additional morphogenetic proteins well documented in B. subtilis, such as CotH and

CotO (Figure 28) [START_REF] Naclerio | Bacillus subtilis spore coat assembly requires cotH gene expression[END_REF], Zilhao et al. 1999[START_REF] Mcpherson | Characterization of the Bacillus subtilis spore morphogenetic coat protein CotO[END_REF], Kim et al. 2006). Both proteins interact with CotE, and their absence results in a disorganized outer coat structure. However, unlike CotE, they do not appear to play a role in attachement of the coat to the FS [START_REF] Zilhao | Assembly requirements and role of CotH during spore coat formation in Bacillus subtilis[END_REF][START_REF] Little | Functional analysis of the Bacillus subtilis morphogenetic spore coat protein CotE[END_REF][START_REF] Mcpherson | Characterization of the Bacillus subtilis spore morphogenetic coat protein CotO[END_REF].

B. subtilis spores do not contain an exosporium. However, analysis using ruthenium red, which stains mainly acid polysaccharides, revealed the presence of filaments on the surface of B. subtilis spores [START_REF] Waller | Ruthenium red staining for ultrastructural visualization of a glycoprotein layer surrounding the spore of Bacillus anthracis and Bacillus subtilis[END_REF]). Since polysaccharides and proteins have been detected in spore surface extracts, these filaments are probably glycoprotein in nature.

Therefore, this layer intimately connected to the outer coat and hardly observable by TEM, was named the spore crust (Figure 28) [START_REF] Chada | Morphogenesis of bacillus spore surfaces[END_REF][START_REF] Waller | Ruthenium red staining for ultrastructural visualization of a glycoprotein layer surrounding the spore of Bacillus anthracis and Bacillus subtilis[END_REF][START_REF] Mckenney | A distance-weighted interaction map reveals a previously uncharacterized layer of the Bacillus subtilis spore coat[END_REF]. Three proteins, CotX, CotY and CotZ were shown to play a morphogenetic role in the assembly of the crust and are recruited in a CotE-dependent manner (Figure 28) [START_REF] Mckenney | A distance-weighted interaction map reveals a previously uncharacterized layer of the Bacillus subtilis spore coat[END_REF][START_REF] Imamura | Proteins involved in formation of the outermost layer of Bacillus subtilis spores[END_REF]. CotY and CotZ are cysteine-rich proteins that were shown to directly interact and may be subjected to cross-linking by formation of disulfide bonds [START_REF] Zhang | Cloning and characterization of a cluster of genes encoding polypeptides present in the insoluble fraction of the spore coat of Bacillus subtilis[END_REF], Krajcikova et al. 2009). The function of the spore crust is not yet clear since mutants without spore crust are resistant to heat and lysozyme treatment [START_REF] Zhang | Cloning and characterization of a cluster of genes encoding polypeptides present in the insoluble fraction of the spore coat of Bacillus subtilis[END_REF]). However, cotXYZ mutant spores have an unusual outer coat that is readily disrupted during fixation, embedding, or sectioning for electron microscopy suggesting that the spore crust might contribute to the tight packing of the outer coat [START_REF] Zhang | Cloning and characterization of a cluster of genes encoding polypeptides present in the insoluble fraction of the spore coat of Bacillus subtilis[END_REF].

III.2.b. Coat assembly

Fluorescence microscopy using GFP-tagged coat proteins allowed to map genetic dependencies for the assembly of the coat and to define a network of interactions [START_REF] Kim | The Bacillus subtilis spore coat protein interaction network[END_REF][START_REF] Mckenney | A distance-weighted interaction map reveals a previously uncharacterized layer of the Bacillus subtilis spore coat[END_REF]. The coat proteins fused to GFP whose localization is dependent on SafA (i.e., CwlJ, CotD, CotT, YaaH, YuzC and OxdD), were assigned to the inner coat while those whose localization is dependent on CotE (CotA, CotM, CotS, YncD and YtxO)

were thought to reside in the outer layer (Figure 28). An approximately equal number of coat proteins were assigned to the inner and outer layers [START_REF] Imamura | Localization of proteins to different layers and regions of Bacillus subtilis spore coats[END_REF][START_REF] Mckenney | A distance-weighted interaction map reveals a previously uncharacterized layer of the Bacillus subtilis spore coat[END_REF]. The subset of CotE-dependent proteins shown to be also dependent on CotXYZ, were therefore categorized as crust proteins (CotW) (Figure 28) [START_REF] Imamura | Localization of proteins to different layers and regions of Bacillus subtilis spore coats[END_REF][START_REF] Mckenney | A distance-weighted interaction map reveals a previously uncharacterized layer of the Bacillus subtilis spore coat[END_REF]. There is also an additional subset of proteins such as YhaX, LipC and YheD, whose localization is dependent on SpoIVA and SpoVM, but independent on both SafA and CotE. The organization of the coat interaction network is highly hierarchical. Each morphogenetic protein is connected to many coat proteins. SpoIVA is at the top of the hierarchy for recruitment of coat proteins, being crucial for the localization of both SafA and CotE on the surface of the developing spore. In contrast, SpoVM and SpoVID are at the top of the spore encasement hierarchy (Figure 28). Encasement of SafA-GFP and CotE-GFP is dependent on SpoVM and SpoVID, and SpoVID interacts directly with SafA and CotE (Figure 28) (Costa et al. 2006, Mullerova et al. 2009, Wang et al. 2009a, de Francesco et al. 2012). The molecular mechanism driving encasement remains unknown but seems to occur in successive coordinated waves, beginning with proteins of the inner coat, followed by the outer coat and finally the crust.

III.2.c. Coat morphogenesis in real time

Apart from the role played by the morphogenetic proteins, transcriptional regulation has important consequences for the production and the dynamic of localization of coat proteins in the MC (Costa et al. 2007). Coat and crust/exosporium assembly takes about 6 h and is mainly governed by the MC. Specific sigma factors and transcription factors creates an hierarchical organization of gene expression that results in successive waves of protein synthesis crucial for proper spore coat assembly [START_REF] Eichenberger | The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis[END_REF], Henriques and Moran 2007, McKenney et al. 2013[START_REF] Driks | The Spore Coat[END_REF]. The major coat morphogenetic proteins are expressed in an appropriate order (Figure 29). The cascade is initiated by σ E that control genes encoding the regulatory proteins SpoIIID and GerR (Figure 21). Later, σ E is replaced by σ K , which regulates GerE-encoding gene (Errington 2003, Piggot andHilbert 2004). The combination of σ E with the regulators SpoIIID and GerR, and of σ K with the regulator GerE, produces a series of coherent and incoherent feedforward loops that result in different temporal patterns (prolonged, delayed, pulsed) of expression for different sets of genes (Figure 29) [START_REF] Eichenberger | The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis[END_REF]). For instance, SpoIIID regulates some genes of the σ E regulon, both positively and negatively, resulting in three waves of gene expression: the first wave is σ E -dependent and will be later repressed by SpoIIID including spoIVA whose product (SpoIVA) is known to finish encasement at the end of engulfment. The second wave is also under the control of σ E alone but unaffected by SpoIIID (including safA),

and the third wave that is σ E -dependent and further activated by SpoIIID [START_REF] Roels | Characterization of spoIVA, a sporulation gene involved in coat morphogenesis in Bacillus subtilis[END_REF][START_REF] Price | A four-dimensional view of assembly of a morphogenetic protein during sporulation in Bacillus subtilis[END_REF], Takamatsu et al. 1999[START_REF] Eichenberger | The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis[END_REF]). In the case of the crust protein operon, cotYZ is initially transcribed at a modest level under the control of σ E , and then transcribed at a high level during late sporulation under the dual control of σ K and GerE [START_REF] Zhang | Cloning and characterization of a cluster of genes encoding polypeptides present in the insoluble fraction of the spore coat of Bacillus subtilis[END_REF][START_REF] Eichenberger | The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis[END_REF]). The encasement by CotZ begins only after the spore becomes visible by phase contrast microscopy. The timing of gene expression is crucial and critical to spore coat morphology. For example, cotE is expressed in two pulses, first from a σ E -dependent promoter that is shut off by SpoIIID and then from a promoter recognized by σ K and is repressed at later stages by GerE (Zheng et al. 1988[START_REF] Eichenberger | The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis[END_REF]). However, a delayed expression of this gene produced spores on which CotE, but not other proteins of the outer layer, was assembled (Costa et al. 2007).

In summary, SpoIVA is recruited to the FS outer membrane by SpoVM (Figure 28).

SpoIVA then recruits SpoVID, whose LysM domain at the C-terminal facilitates its binding to the cortex PG beneath the FS outer membrane (Figure 28). SpoIVA, SpoVM and SpoIVD marks the MC proximal pole as the site of coat attachment and the three proteins finish spore encasement as soon as engulfment is completed (Figure 29) (McKenney and Eichenberger 2012). SafA is targeted to the MC proximal pole first in a SpoIVD-independent way, which probably depends on its LysM domain present at the N-terminal (Figure 28). CotE recruitment is most likely mediated by SpoIVA. Therefore, a scaffold organized in multiple layers is assembled on one pole of the FS (Figure 29). Subsequently, the coat encases the spore in multiple coordinated waves. Importantly, additional coat proteins can be added to this scaffold even after completion of encasement. This property implies that the coat remains permeable to proteins of a certain size until late stages in sporulation. For instance, the coat protein CotD is among the last to be produced (class VI), but eventually localizes to the inner

III.3. Spore coat of B. cereus group

Differences in the appearance and thickness of the coat layers among spores of various bacilli strains or species are observed. The B. cereus group includes B. anthracis and B.

thurigiensis. The coat of B. cereus and B. anthracis is thinner and more compact than those of B. subtilis (Figure 30A) [START_REF] Chada | Morphogenesis of bacillus spore surfaces[END_REF]. In B. thuringiensis, the inner coat is laminated but consists of a patchwork of striated packets, appearing either stacked or comb-like, and the outer coat is granular (Figure 30B) [START_REF] Gerhardt | Fine structure of the Baccilus thuringiensis spore[END_REF]). B. subtilis and B. anthracis group have a similar number of coat proteins. However, differences in the electrophoretic patterns suggest that the biochemical composition of the coat is different [START_REF] Kuwana | Proteomics characterization of novel spore proteins of Bacillus subtilis[END_REF][START_REF] Lai | Proteomic analysis of the spore coats of Bacillus subtilis and Bacillus anthracis[END_REF][START_REF] Redmond | Identification of proteins in the exosporium of Bacillus anthracis[END_REF]). Since all bacilli contain the morphogenetic proteins SpoIVA, SpoVM and CotE, and almost all have SpoVID and SafA [START_REF] Lai | Proteomic analysis of the spore coats of Bacillus subtilis and Bacillus anthracis[END_REF], Liu et al. 2004[START_REF] Giorno | Morphogenesis of the Bacillus anthracis spore[END_REF], it is tempting to speculate that nearly all bacilli share the mechanisms of coat assembly. Indeed, B. anthracis and B. subtilis use some of the same proteins to direct assembly of their coat. SpoIVA is necessary to anchor the coat to the FS surface and for cortex formation in both species [START_REF] Roels | Characterization of spoIVA, a sporulation gene involved in coat morphogenesis in Bacillus subtilis[END_REF][START_REF] Giorno | Morphogenesis of the Bacillus anthracis spore[END_REF]). However, in B. anthracis, CotE plays only a modest role in coat formation, but a major role for proper assembly of the outermost layer, the exosporium. In this bacterium, it is rather CotH that directs the assembly of an important subset of coat proteins. Accordingly, cotH deletion has a more severe impact on coat assembly than cotE inactivation [START_REF] Giorno | Morphogenesis of the Bacillus anthracis spore[END_REF]). In B. cereus, exsA is the equivalent of B. subtilis safA and the gene organizations around both are identical.

The location of ExsA has not been determined but it possesses an N-terminal LysM domain, as SafA, although the rest of the sequence is less conserved. ExsA, which controls coat and exosporium assembly in B. cereus, appears to have a more severe impact on the B. cereus coat assembly than does safA in B. subtilis [START_REF] Bailey-Smith | The ExsA protein of Bacillus cereus is required for assembly of coat and exosporium onto the spore surface[END_REF].

III.4. The Exosporium layer

The exosporium is the outermost layer of the B. cereus spores and those of its close relatives B. anthracis and B. thuringensis [START_REF] Stewart | The Exosporium Layer of Bacterial Spores: a Connection to the Environment and the Infected Host[END_REF]. The exosporium is a hydrophobic, balloon-like glycoprotein layer separated from the spore coat by a gap called the interspace (Figure 30) (Henriques and Moran 2007[START_REF] Giorno | Localization and assembly of proteins comprising the outer structures of the Bacillus anthracis spore[END_REF][START_REF] Kailas | Surface architecture of endospores of the Bacillus cereus/anthracis/thuringiensis family at the subnanometer scale[END_REF]. It consists of a paracrystalline basal layer adorned with hair-like projections (Figure 30) (Gerhardt andRibi 1964, Wehrli et al. 1980). In B. cereus group, the exosporium composition and structure are very similar [START_REF] Ball | Structure of the exosporium and sublayers of spores of the Bacillus cereus family revealed by electron crystallography[END_REF] and is anchored to the coat layer via a number of proteinprotein interactions. Proteins implicated in exosporium attachment include CotE, CotY, ExsA, ExsB, ExsY and ExsM (Bailey-Smith et al. 2005[START_REF] Boydston | The ExsY protein is required for complete formation of the exosporium of Bacillus anthracis[END_REF][START_REF] Johnson | ExsY and CotY are required for the correct assembly of the exosporium and spore coat of Bacillus cereus[END_REF][START_REF] Mcpherson | ExsB, an unusually highly phosphorylated protein required for the stable attachment of the exosporium of Bacillus anthracis[END_REF][START_REF] Stewart | The Exosporium Layer of Bacterial Spores: a Connection to the Environment and the Infected Host[END_REF]. The basal layer of the exosporium contains over a dozen different integral proteins including CotY, ExsY, ExsFA and ExsFB [START_REF] Redmond | Identification of proteins in the exosporium of Bacillus anthracis[END_REF][START_REF] Stewart | The Exosporium Layer of Bacterial Spores: a Connection to the Environment and the Infected Host[END_REF]. The hair-like projections are attributed to the presence of the collagenlike glycoprotein, BclA, which is the major component of B. anthracis exosporium and is homogenously distributed at the spore surface (Figure 30A) [START_REF] Sylvestre | A collagen-like surface glycoprotein is a structural component of the Bacillus anthracis exosporium[END_REF]. The Nterminus of BclA is covalently attached to the basal layer while the C-terminus of BclA is oriented to the exosporium surface [START_REF] Boydston | Orientation within the exosporium and structural stability of the collagen-like glycoprotein BclA of Bacillus anthracis[END_REF]. Localization of B. anthracis BclA is mediated in part by the basal proteins ExsFA, ExsFB proteins [START_REF] Sylvestre | Contribution of ExsFA and ExsFB proteins to the localization of BclA on the spore surface and to the stability of the bacillus anthracis exosporium[END_REF][START_REF] Thompson | The co-dependence of BxpB/ExsFA and BclA for proper incorporation into the exosporium of Bacillus anthracis[END_REF]. As for the B. subtilis spore crust, cysteine-rich proteins are also essential for the assembly of the exosporium. ExsY and CotY of B. anthracis and B. cereus are orthologues of the B. subtilis CotZ protein [START_REF] Boydston | The ExsY protein is required for complete formation of the exosporium of Bacillus anthracis[END_REF][START_REF] Johnson | ExsY and CotY are required for the correct assembly of the exosporium and spore coat of Bacillus cereus[END_REF][START_REF] Imamura | Proteins involved in formation of the outermost layer of Bacillus subtilis spores[END_REF]. Whether the crust is functionally equivalent to the surface of the exosporium and whether crust-like layers are found on the outer spore coat in exosporiumcontaining spores are questions that remains unanswered. Some studies have proposed that the role of the exosporium is to protect the spore against degradative enzymes, antibodies, and macrophages (NO), while others suggest that the exosporium helps to regulate spore germination and adherence to abiotic and biotic surfaces [START_REF] Tauveron | Variability among Bacillus cereus strains in spore surface properties and influence on their ability to contaminate food surface equipment[END_REF][START_REF] Weaver | Protective role of Bacillus anthracis exosporium in macrophage-mediated killing by nitric oxide[END_REF][START_REF] Faille | Morphology and physico-chemical properties of Bacillus spores surrounded or not with an exosporium: consequences on their ability to adhere to stainless steel[END_REF][START_REF] Stewart | The Exosporium Layer of Bacterial Spores: a Connection to the Environment and the Infected Host[END_REF]. Present in the exosporium layer, an Alanine racemase that converts L-alanine (a germinant for B. anthracis) to D-alanine (an inhibitor of germination) may function to inhibit premature germination under suboptimal growth conditions [START_REF] Chesnokova | The sporespecific alanine racemase of Bacillus anthracis and its role in suppressing germination during spore development[END_REF]). An Arginase activity is also present at the surface of B. anthracis spores [START_REF] Raines | Importance of nitric oxide synthase in the control of infection by Bacillus anthracis[END_REF], which could compete with the macrophage nitric oxide synthase (NOS2) that use L-arginine substrate to produce NO [START_REF] Weaver | Protective role of Bacillus anthracis exosporium in macrophage-mediated killing by nitric oxide[END_REF]). In addition, two spore SODs, Sod15 and SodA1, are present in the exosporium and their activity can afford B. anthracis protection against oxidative stress [START_REF] Cybulski | Four superoxide dismutases contribute to Bacillus anthracis virulence and provide spores with redundant protection from oxidative stress[END_REF]. While interactions between the exosporium protein BclA and the host integrin Mac-1 (CR3) allow entry of B. anthracis spores into phagocytic host cells [START_REF] Oliva | The integrin Mac-1 (CR3) mediates internalization and directs Bacillus anthracis spores into professional phagocytes[END_REF], the exosporium protects the spores from macrophage-mediated killing [START_REF] Weaver | Protective role of Bacillus anthracis exosporium in macrophage-mediated killing by nitric oxide[END_REF]. Finally, the metalloprotease InhA1 present in the exosporium of B. cereus allow bacterium to escape from macrophages [START_REF] Ramarao | The InhA1 metalloprotease allows spores of the B. cereus group to escape macrophages[END_REF].

A ballon-like exosporium has been also observed in clostridia such as Clostridium sporogenes and Clostridium sordellii. However, very different structure and protein compositions were identified when compared to B. cereus group [START_REF] Janganan | Characterization of the spore surface and exosporium proteins of Clostridium sporogenes; implications for Clostridium botulinum group I strains[END_REF][START_REF] Rabi | Clostridium sordellii outer spore proteins maintain spore structural integrity and promote bacterial clearance from the gastrointestinal tract[END_REF]. C. sordelli exosporium is composed by a thick inner layer and a thin outer layer, do not possess hair-like projections and is open at the spore poles [START_REF] Rabi | Structural Characterization of Clostridium sordellii Spores of Diverse Human, Animal, and Environmental Origin and Comparison to Clostridium difficile Spores[END_REF].

Apertures in the exosporium and a balloon-like exosporium seem to be a characteristic of clostridial spores. Indeed, an opening in the exosporium has also been reported in C. sporogenes and C. botulinum [START_REF] Stevenson | Variability in ultrastructure of Clostridium botulinum spores[END_REF][START_REF] Brunt | Apertures in the Clostridium sporogenes spore coat and exosporium align to facilitate emergence of the vegetative cell[END_REF][START_REF] Rabi | Structural Characterization of Clostridium sordellii Spores of Diverse Human, Animal, and Environmental Origin and Comparison to Clostridium difficile Spores[END_REF]. The function of an open exosporium remains unclear but it very likely allows entry of larger molecules into the exosporium rather than acting as a protective barrier. In C. sporogenes, newly formed vegetative bacteria were shown to emerge through the pre-existing aperture in the exosporium openings upon germination [START_REF] Brunt | Apertures in the Clostridium sporogenes spore coat and exosporium align to facilitate emergence of the vegetative cell[END_REF].

III.5. Spore surface layers of C. difficile

TEM studies indicate that the coat of C. difficile comprises two layers, consisting of an inner lamellar coat and a more electrodense outer coat (Figure 27C) (Lawley et al. 2009b, Permpoonpattana et al. 2011b, Paredes-Sabja et al. 2012[START_REF] Rabi | Structural Characterization of Clostridium sordellii Spores of Diverse Human, Animal, and Environmental Origin and Comparison to Clostridium difficile Spores[END_REF]. The stability of the exosporium was for a long time a matter of controversy. Several reports suggest that this layer is fragile and easily lost (Permpoonpattana et al. 2011b, Permpoonpattana et al. 2013), whereas other reports indicate that it is a stable layer that is only removed by proteases and/or sonication treatments [START_REF] Joshi | Contribution of spores to the ability of Clostridium difficile to adhere to surfaces[END_REF], Barra-Carrasco et al. 2013, Escobar-Cortes et al. 2013, Pizarro-Guajardo et al. 2014). Nowadays, it become clear that the stability of this layer could be related with the strain or to proteases used during spore purification procedures (Escobar-Cortes et al. 2013[START_REF] Pizarro-Guajardo | Ultrastructural Variability of the Exosporium Layer of Clostridium difficile Spores[END_REF]. C. difficile exosporium is a proteinaceous layer that contributes to the hydrophobicity of the spore surface (Diaz-Gonzalez et al. 2015). In contrast to the spore coat, the exosporium can be removed by enzymatic treatment with proteases (i.e., trypsin or proteinase K) or by sonication (Escobar-Cortes et al. 2013). Several studies showed that the morphology of the exosporium is straindependent [START_REF] Joshi | Contribution of spores to the ability of Clostridium difficile to adhere to surfaces[END_REF], Escobar-Cortes et al. 2013[START_REF] Pizarro-Guajardo | Ultrastructural Variability of the Exosporium Layer of Clostridium difficile Spores[END_REF].

Whereas spores of the 630 strain have an electron-dense compact exosporium layer, in most of others C. difficile strains studied (R20291, M120, TL176, and TL178), the exosporium layer have hair-like extensions similar as those observed in spore of the B. cereus group [START_REF] Chada | Morphogenesis of bacillus spore surfaces[END_REF], Paredes-Sabja et al. 2014[START_REF] Pizarro-Guajardo | Ultrastructural Variability of the Exosporium Layer of Clostridium difficile Spores[END_REF]. In contrast to other spore formers, during sporulation C. difficile forms two distint exosporium morphotypes [START_REF] Pizarro-Guajardo | Ultrastructural Variability of the Exosporium Layer of Clostridium difficile Spores[END_REF]). These morphotypes include either spores with a thick or a thinner-exosporium layer as determined by the thickness of the electrodense material surrounding the coat [START_REF] Pizarro-Guajardo | Ultrastructural Variability of the Exosporium Layer of Clostridium difficile Spores[END_REF]. Notably, the exosporium layer of C. difficile spores differs from that of spores of the B. cereus group [START_REF] Stewart | The Exosporium Layer of Bacterial Spores: a Connection to the Environment and the Infected Host[END_REF][START_REF] Pizarro-Guajardo | Ultrastructural Variability of the Exosporium Layer of Clostridium difficile Spores[END_REF][START_REF] Rabi | Structural Characterization of Clostridium sordellii Spores of Diverse Human, Animal, and Environmental Origin and Comparison to Clostridium difficile Spores[END_REF]. In fact, C. difficile lacks many exosporium proteins of the B. cereus group's spores as well as the frequently observed gap that separates the spore coat from the exosporium. Instead, the exosporium of C. difficile spores is in direct contact with the spore coat layers in a similar way as the crust of B. subtilis spores (Barra-Carrasco et al. 2013, Paredes-Sabja et al. 2014, Diaz-Gonzalez et al. 2015, Calderon-Romero et al. 2018).

III.5.a. Morphogenesis of the C. difficile spore coat

Only 25% of the B. subtilis spore coat proteins have homologs in C. difficile and most of the morphogenetic proteins identified in B. subtilis or other bacilli are absent in C. difficile.

This suggests that the components that govern spore coat assembly in C. difficile are divergent to those defined for B. subtilis and/or the B. cereus group (Henriques andMoran 2007, Lawley et al. 2009b). Among spore coat morphogenetic proteins, only SpoIVA is present in all bacilli and clostridia (Galperin et al. 2012). Since SpoVM is very short (less than 30 aa), it is difficult to identify its encoding gene from genome annotation. In addition, the absence of SpoIVD, SafA and CotE in clostridia suggests major differences in the assembly of the bacilli and clostridia surface layers (Galperin et al. 2012). In C. difficile, homologs for SpoIVA and SpoVM have been identified (Galperin et al. 2012). It is reasonable to propose that the SpoVM-SpoIVA-dependent mechanism for attaching the coat to the FS outer membrane was common in the two groups of spore forming bacteria.

Consistent with this, C. difficile SpoIVA was shown to be essential for coat localization around the FS as observed in B. subtilis (Putnam et al. 2013). Indeed, in a spoIVA mutant, the coat is mislocalized and detected in the MC cytoplasm as in B. subtilis, where distinct coat layers are observed in the mislocalized coat swirls. This suggested that coat can still assemble, but SpoIVA is required to localize the coat around the FS. However, in contrast to B. subtilis, C. difficile SpoIVA is dispensable for cortex synthesis [START_REF] Roels | Characterization of spoIVA, a sporulation gene involved in coat morphogenesis in Bacillus subtilis[END_REF], Putnam et al. 2013).

The spoVM gene of C. difficile encodes a small protein that like in B. subtilis is expressed in the MC under the control of σ E . Moreover, many of the residues previously identified to be critical for the function of B. subtilis SpoVM are conserved in SpoVM of C. difficile.

However, contrary to what we observed in B. subtilis, C. difficile SpoVM was found to be dispensable for spore formation/maturation (Ribis et al. 2017). The inactivation of spoVM leads to morphological defects in a small percentage of cells, such as coat detachment from the FS or abnormal cortex thickness. It has been shown that C. difficile SpoVM and SpoIVA directly interact as observed in B. subtilis. However, SpoIVA is still able to polymerize on the surface of FS in the spoVM mutant and to encase the FS (Ribis et al. 2017). Even if the the spore coat layers of B. subtilis and C. difficile are morphologically similar, the absence of SpoVID, SafA and CotE homologues in C. difficile, suggest that different proteins replace these morphogenetic proteins involved in the spore coat assembly. A functional homolog of B. subtilis SpoVID (CD3567) has been identified in C. difficile. This protein, contains a C-LysM domain (like SpoVID) that was shown to directly interact with the Walker A ATPbinding motif of SpoIVA, suggesting that CD3567 and SpoIVA form a complex that may drive coat assembly around the FS (Putnam et al. 2013). CD3567 was then renamed SipL for "SpoIVA interacting protein L". However, the mechanism by which SpoIVA recognizes SipL fundamentally differs from that of SpoIVA recognizing SpoVID in B. subtilis. In addition, a sipL mutant shows a phenotype reminescent of the spoIVA mutant, where the spore coat is tethered and mislocalized to the MC cytoplasm (Putnam et al. 2013). SipL, as SpoVID, might play a role in the encasement step, helping in the transition of the coat from the MC proximal pole (MCP) to full spherical shell around the spore (Wang et al. 2009a, McKenney andEichenberger 2012). Whether the LysM domain tethers SipL to the FS by interacting with the cortex in addition to its presumed interaction with SpoIVA is still unkown.

III.5.b. Spore coat and exosporium proteins

Proteomic studies allow identifying spore proteins localized at the surface layers of C. difficile spores i.e., coat and exosporium (Permpoonpattana et al. 2011b, Abhyankar et al. 2013, Permpoonpattana et al. 2013, Diaz-Gonzalez et al. 2015). For several proteins, their presence at the spore surface was confirmed using antibodies raised against them either by immunofluorescence or by Western blot. CotA (CD1613) and CotB (CD1511) were identified at the surface of C. difficile spore. The cotA mutant spores exhibited major morphological defects and lacked the outer coat layer for half of them and were more sensitive to lysozyme (Permpoonpattana et al. 2013). By contrast, no detectable differences in spore ultrastructure or lysozyme resistance were observed for the cotB mutant spores. CotF (CD0597) is a tyrosine-rich protein homolog to B. subtilis CotJB, likely a structural component of the inner layers of the coat [START_REF] Henriques | Characterization of cotJ, a sigma E-controlled operon affecting the polypeptide composition of the coat of Bacillus subtilis spores[END_REF][START_REF] Seyler | Assembly and interactions of cotJ-encoded proteins, constituents of the inner layers of the Bacillus subtilis spore coat[END_REF]. CotF is located at the spore surface but its role remains to be investigated (Permpoonpattana et al. 2013). CotCB (CD0598), CotD (CD2401), and CotG (CD1567) are a group of proteins predicted to have Mn 2+ catalase activity (Permpoonpattana et al. 2011b, Permpoonpattana et al. 2013). Indeed, catalase activity was observed for the recombinant proteins for CotD and CotG while no activity was found for CotCB (Permpoonpattana et al. 2013). CotCB and CotD are homologs to the B. subtilis outer coat protein CotJC [START_REF] Henriques | Characterization of cotJ, a sigma E-controlled operon affecting the polypeptide composition of the coat of Bacillus subtilis spores[END_REF][START_REF] Seyler | Assembly and interactions of cotJ-encoded proteins, constituents of the inner layers of the Bacillus subtilis spore coat[END_REF]. As for cotJC mutant of B. subtilis [START_REF] Henriques | Characterization of cotJ, a sigma E-controlled operon affecting the polypeptide composition of the coat of Bacillus subtilis spores[END_REF], no ultrastructural defects were observed in cotD and cotCB mutant spores (Permpoonpattana et al. 2013). Other proteins with enzymatic activities are present at the surface of C. difficile spores, such as SodA (CD1631), a homolog of the B. subtilis superoxide dismutase (SodA) and CotE, a bifunctional protein with peroxiredoxin reductase (N-terminal) and chitinase (C-terminal) activities. Even if the recombinant SodA protein carries high levels of SOD activity, the loss of C. difficile sodA had no effect on spore ultrastructure or spore resistance (Permpoonpattana et al. 2013). There are also no obvious ultrastructural defects in spores of a cotE mutant (Permpoonpattana et al. 2011b, Permpoonpattana et al. 2013). Chitinase and a low level of peroxidase activities were detected with recombinant CotE C and CotE N , respectively (Permpoonpattana et al. 2013).

Interestingly, it was found recently that CotE plays an important role in spore attachment to the gut mucosa via direct interaction with mucin and the chitinase activity of CotE contributes to mucin degradation [START_REF] Hong | The Spore Coat Protein CotE Facilitates Host Colonization by Clostridium difficile[END_REF] CotA, CotB, CotD and CotE are among the proteins that were identified in the exosporium extracts (Diaz-Gonzalez et al. 2015). However, analysis of untreated and proteinase K-treated spores revealed that CotA and CotB are surface exposed and accessible to antibodies but that most of these proteins are insentive to proteinase K treatment confirming their localization in the spore coat (Diaz-Gonzalez et al. 2015). BclA proteins, as well as the cysteine-rich proteins CdeC, CdeM and CdeA are mainly located in the exosporium layer since these proteins are almost absent from the surface of proteinase K-treated spores (Diaz-Gonzalez et al. 2015). In [START_REF] Sebaihia | The multidrug-resistant human pathogen Clostridium difficile has a highly mobile, mosaic genome[END_REF]. The inactivation of bclA1 and bclA2 genes causes aberrations in the spore coat, and spores of all three independent bclA mutants showed reduced surface hydrophobicity and a faster germination rate [START_REF] Phetcharaburanin | The spore-associated protein BclA1 affects the susceptibility of animals to colonization and infection by Clostridium difficile[END_REF]. Glycosylation of the C. difficile BclA3 protein at the spore surface have been demonstrated. This is consistent with the presence of the sgtA gene located immediately upstream of the blclA3, which encodes a glycosyltransferase involved in the biosynthesis of surface-associated glycan components [START_REF] Strong | Identification and characterization of glycoproteins on the spore surface of Clostridium difficile[END_REF].

Unlike 630 spores, R20291 and CD196 strains have a pseudogene bclA1 due to the appearance of an early stop codon. As consequence, the 027 strains encode a BclA1 protein of approximately 6 kDa that still can assemble at the spore surface [START_REF] Stabler | Comparative genome and phenotypic analysis of Clostridium difficile 027 strains provides insight into the evolution of a hypervirulent bacterium[END_REF][START_REF] Phetcharaburanin | The spore-associated protein BclA1 affects the susceptibility of animals to colonization and infection by Clostridium difficile[END_REF].

III.5.c. Morphogenetic proteins of C. difficile exosporium

As in bacilli, cysteine-rich proteins are also present at the surface of C. difficile spores.

A previous study performed in R20291 strain, demonstrated that the C. difficile exosporium cysteine-rich protein "CdeC" (CD1067), is required for exosporium morphogenesis and coat assembly (Barra-Carrasco et al. 2013). Very recently, new insights into the mechanism of C. difficile exosporium assembly were brought with the study of CdeC and CdeM in the 630 strain (Calderon-Romero et al. 2018). cdeC and cdeM are transcribed by σ K . While cdeC is conserved in related Peptostreptococcaeace family members, cdeM is unique to C. difficile.

These two cysteine-rich proteins have different role in the integrity of the spore surface layers. Both proteins are required for the correct formation of the exosporium layer. However,

Objectives of the thesis

During its infectious cycle, C. difficile is exposed to various stresses. In the environment, spores face atmospheric O 2 and disinfectants used for their eradication. Then, inside the host both spores and vegetative cells are exposed, among other stresses, to O 2 and reactive oxygen species produced by the host cells during inflammation. Therefore, C.

difficile must be able to protect itself and survive in these hostile environments. The spore surface layers typically enable spore survival in the presence of predatory enzymes and chemical compounds found in the gastrointestinal tract. However, even if some progress has been made in understanding the biology of C. difficile spore formation, many questions remain about morphogenesis of the spore surface layers and the mechanisms used by spores to protect against toxic compounds. In the gut, once spore germination is complete, vegetative cells need to protect against low O 2 tensions as well as oxidative and nitrosative conditions.

However, little is still known about the proteins involved in the ability of C. difficile to tolerate O 2 , ROS and RNS. C. difficile genome encodes both revRbrs and FDPs, suggesting the existence of O 2 and ROS detoxification pathways in this bacterium. Interestingly, some of these proteins were detected in the proteome of the spore surface layers, which suggests that such detoxification can also be done by spores.

The first objective of my thesis is to study some members of the σ K -regulon, including both CD1065 and CD2399 genes, and their role in the spore formation of C. difficile.

The second objective is 1) to study the expression of the genes encoding the revRbrs and FDPs during both vegetative growth and sporulation; 2) to characterize the enzymatic reactions catalyzed by some of them; and 3) to study the role of these proteins in the management of C. difficile O 2 tolerance.

Results

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Spore coat morphogenesis Nicholson 2000, [START_REF] Klobutcher | The Bacillus subtilis spore coat provides "eat resistance" during phagocytic predation by the protozoan Tetrahymena thermophila[END_REF][START_REF] Setlow | Spore Resistance Properties[END_REF]. Additionally, the spore surface is likely the primary site of spore-host interactions [START_REF] Stewart | The Exosporium Layer of Bacterial Spores: a Connection to the Environment and the Infected Host[END_REF]. Therefore, understanding fundamental aspects on morphogenesis of the spore surface layers and their functional properties is a matter of great interest.

The structure of C. difficile spore is similar to that of the other spore-forming bacteria.

However, the protein composition of the spore surface layers differs from that of bacilli To make the reading easier, supplementary tables S1, S3 and S4 from this article are in Annex. Due to the large amount of raw data, the table S2 concerning the comparative analysis of the spore coat/exosporium proteome from the wild-type and the cotL mutant is not included in paper format in the manuscript. The supplementary figures S1, S2, S3, S4, S5 and S6 are in the text after all the main figures.

Introduction

Clostridioides difficile, commonly known as Clostridium difficile, is a Gram-positive, spore-forming obligate anaerobic bacterium that is currently considered to be the leading cause of antibiotic-associated diarrhoea (Kelly andLaMont 2008, Rupnik et al. 2009). The incidence and severity of C. difficile infection (CDI) have been increasing in North America and Europe (Rupnik et al. 2009, Wiegand et al. 2012). Spores, which are the primary infectious form, play a crucial role in transmission, infection and recurrence (Lawley et al. 2010, Deakin et al. 2012). As an obligate anaerobic bacterium, vegetative cells are unable to grow in an aerobic environment, and the metabolically dormant spore has a central role in the resistance outside the host. After ingestion, the earliest CDI events correspond to spore germination upon sensing specific bile salts in the gastrointestinal tract, followed by the growth of vegetative cells and gut colonization (Sorg and Sonenshein 2008). The vegetative cells then produce two toxins, TcdA and TcdB, which are the primary virulence factors required for the development of CDI symptoms. Spores are also produced in the gut and cause both spreading into the environment and relapse (Deakin et al. 2012, Janoir et al. 2013). The resistance to extreme environmental conditions and chemical compounds, including oxygen, antibiotics and disinfectants, makes the spores difficult to eradicate (Rupnik et al. 2009, Edwards et al. 2016a).

The process of sporulation is an ancient bacterial developmental programme that has been extensively studied in the model organism Bacillus subtilis (Hilbert andPiggot 2004, Higgins andDworkin 2012). Spore morphogenesis starts with the formation of an asymmetric septum, which produces a smaller forespore (FS) and a larger mother cell (MC). The MC then engulfs the FS and directs the formation of the spore protective layers that are the cortex, coat and exosporium (Henriques andMoran 2007, McKenney andEichenberger 2012). After a period of spore maturation, the MC undergoes autolysis to release the spore. The programmes of gene expression in the FS and MC compartments are defined by the cell type-specific RNA polymerase sigma (σ) factors: σ F in the FS and σ E in the MC control the early stages of sporulation and are replaced following engulfment completion by σ G and σ K in the respective compartment (Galperin et al. 2012, Abecasis et al. 2013). In C. difficile, as observed in B.

subtilis, σ F and σ G activity is restricted to the FS, and σ E and σ K govern development events in the MC (Pereira et al. 2013, Saujet et al. 2013). Interestingly, significant deviations from the B. subtilis paradigm concerning the regulatory circuit leading to the σ factor activation are observed with a reduced communication between the MC and the FS and a weaker connection between gene expression and morphogenesis (Pereira et al. 2013, Saujet et al. 2013). C.

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difficile spore structure is similar to that of other spore-forming bacteria. From the internal to external side, the spores are comprised of a dehydrated central core surrounded by the inner FS membrane, a germ cell wall, a modified peptidoglycan cortex layer, an outer membrane, a proteinaceous layer known as the spore coat and an outermost layer named the exosporium (Henriques andMoran 2007, Paredes-Sabja et al. 2014). However, the protein composition of the spore surface layers in C. difficile differs from that of other bacilli and clostridia (Lawley et al. 2009b, Permpoonpattana et al. 2011b, Abhyankar et In C. difficile, the absence of homologs for B. subtilis SafA and CotE suggests that different proteins are involved in the assembly of the spore surface layers (Galperin et al. 2012).

In C. difficile, as in B. subtilis, the spore surface layers are comprised of proteins produced by the MC under the control of σ E and/or σ K (Pereira et al. 2013, Saujet et al. 2013, Pishdadian et al. 2015). In a previous study, transcriptome analyses combined with a genome-wide transcriptional start site mapping allowed the definition of the regulons of the four σ factors of sporulation, σ F , σ E , σ G and σ K in C. difficile (Saujet et al. 2013). Among the genes whose expression is controlled by the MC specific σ factors, we identified CD1065 encoding a lysine-rich protein of unknown function with no homologs in bacilli. The expression of the CD1065 gene is down-regulated in a sigE and sigK mutant (Fimlaid et al. 2013, Saujet et al. 2013) and is induced during the infection of axenic mice (Janoir et al. 2013). In this study, we showed that the CD1065 gene is transcribed by σ E and σ K during endospore development. We determined that the CD1065 mutant forms spores lacking most of the spore coat and exosporium proteins, while the level of gene expression or protein production of the coat/exosporium components was similar in the sporulating cells of the CD1065 mutant and the wild-type strain. In addition, CD1065 spores are sensitive to lysozyme and impaired in germination. These studies characterize a new coat protein crucial for the assembly of the spore surface layers in C. difficile and provide new information about the assembly partners of the spore coat in this bacterium.

Results

Expression of CD1065 in the mother cell compartment is under the dual control of the sporulation sigma factors σ E and σ K

In C. difficile, σ K is essential for the assembly of the spore surface layers (Fimlaid et al. 2013, Pereira et al. 2013, Saujet et al. 2013). In the genome-wide analysis of cell typespecific transcription during spore formation, CD1065 appears as a member of the σ K regulon (Saujet et al. 2013). As expected, the expression of CD1065 decreases 17-and 3-fold in the sigE and sigK mutants, respectively, in qRT-PCR experiments (Saujet et al. 2013). In this study, we identified two transcriptional start sites (TSSs) located 116 nt (promoter P1) and 42 nt (promoter P2) upstream of the translational start site of CD1065 (Fig. 1A). A canonical -10 box (CATATATA) for promoters recognized by σ E is present upstream of the TSS corresponding to P1, and an ATT sequence 14 bp upstream from the -10 box could match the -35 motif. Upstream of the second TSS (P2), we found a -10 (TATATAAA) and a -35 (AC)

box close to the consensus of the σ K -dependent promoters (Fig. 1A).

The σ E -and σ K -dependent promoters identified upstream of CD1065 suggested that this gene might be expressed in the MC compartment from the early to late stages of sporulation as demonstrated for sigK expression itself (Pereira et al. 2013, Serrano et al. 2016b). To monitor the CD1065 expression during sporulation, the promoter region containing both the σ E -and the σ K -dependent promoters was cloned into pFT47, a vector carrying the gene encoding a fluorescent reporter SNAP Cd (Pereira et al. 2013, Cassona et al. 2016). After 24 h of growth in sporulation medium (SM), cells expressing the transcriptional P σE/σK(CD1065) -SNAP Cd fusion were doubly labeled with TMR-Star and the membrane dye MTG to allow identification of the different stages of sporulation. Fluorescence microscopy revealed that the expression of the P σE/σK(CD1065) -SNAP Cd fusion was confined to the MC (Fig. 1B). CD1065 expression was detected immediately after asymmetric division in 50% of the cells, increasing to 68% of the cells during engulfment and to 92% of the cells after engulfment completion. No CD1065 expression was observed in the FS. To investigate the impact of each σ factor on CD1065 expression, we introduced the transcriptional fusion into sigE and sigK mutants. The expression of P σE/σK(CD1065) -SNAP Cd was abolished in the sigE mutant (Fig. 1B). In contrast, sigK inactivation did not suppress the expression of this fusion, since fluorescence was detected similarly to the wild-type 630Δerm strain after engulfment.

When we monitored the SNAP production by a Western blot using an anti-SNAP antibody, no signal was detected in the sigE background (Fig. S1A). However, a decrease in the Results

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abundance of the SNAP protein was observed in the sigK mutant compared to the wild-type strain (Fig. S1A). This result was consistent with the decreased level of the CD1065 transcript detected by qRT-PCR in the sigK mutant (Saujet et al. 2013) even if we failed to detect this decrease by analysing the P σE/σK(CD1065) -SNAP Cd fusion using microscopy. To study the respective roles of σ E and σ K in the complex pattern of expression of CD1065 in more detail, we constructed two additional transcriptional fusions with the individual promoters:

P σE(CD1065) -SNAP Cd and P σK(CD1065) -SNAP Cd . Fluorescence expression from the P σE(CD1065) -SNAP Cd fusion was similar to that observed for the P σE/σK(CD1065) -SNAP Cd fusion, and a signal was only detected in the sigK mutant sporulating cells (Fig. 1C). Regarding the P σK(CD1065) -SNAP Cd fusion, we detected a significantly lower fluorescent signal at the initial stage of morphogenesis with 21% of cells detected after asymmetric division instead of the 50% for the P σE/σK(CD1065) -SNAP Cd fusion (Fig. 1D). Since it is known that σ E is active earlier than σ K , the decreased number of cells expressing fluorescence immediately after asymmetric division might be a result of the delayed expression of the SNAP from the P σK(CD1065) -SNAP Cd fusion when compared to the P σE/σK(CD1065) -SNAP Cd fusion (Pereira et al. 2013, Saujet et al. 2013).

No signal was detected in both the sigE and sigK mutants with the P σK(CD1065) -SNAP Cd fusion (Fig. 1D), which is consistent with the requirement of σ E for sigK expression (Pereira et al. 2013, Saujet et al. 2013). Samples of cultures expressing each fusion collected after 24 h of growth in SM were also analysed using Western blot. A reduced amount of the SNAP protein was detected when the gene is controlled by either σ E or σ K (Fig. S1B) suggesting that both σ factors contribute to CD1065 expression. Together, these results strongly suggested that CD1065 transcription is initiated from P1 by σ E at early stage of sporulation and is maintained or increased during the late stages by the use of both P1 and P 2 recognized by σ E and σ K , respectively.

CD1065 is required for efficient germination

To test whether CD1065 plays a role in sporulation and/or germination, we inserted a group II intron into the CD1065 gene in the 630Δerm strain in a sense orientation immediately after the 123rd nucleotide in its coding sequence (Fig. S2) (Heap et al. 2007, Heap et al. 2009). We cloned the CD1065 gene expressed from both P1 and P2 into the plasmid pMTL84121 for complementation studies. After 24 h of growth in SM, phasecontrast microscopy analysis revealed that the mutant can sporulate. The frequency of spore formation was 20% of the sporulating cells for the mutant compared with 18% for the wildtype strain (Fig. 2A). We purified the spores to analyse their morphology. While the wild-type strain forms only phase-bright spores, the CD1065 mutant produces both mature phase-bright spores and a small proportion (8%) of phase-grey immature spores (Fig. 2B, red arrow). In addition, mutant spores appeared smaller than the wild-type spores. Measurements of the length and width of the spores showed that the mutant spores are thinner and shorter (Fig. 2C). After 24 h of growth in SM, the titre of the heat resistant spores was also assessed by plating them in BHI containing 0.1% of the germinant taurocholate (TA). The CD1065 mutant produced 10-fold fewer heat resistant spores than the wild-type strain.

Complementation partially restored the heat resistant spore formation (Fig. 2D). Since the CD1065 mutant can sporulate, the reduction in heat resistant spores detection might be due to a defect in germination. The addition of a germinant, such as TA, results in a change from refractive phase-bright spores to phase-dark spores due to cortex hydrolysis, calcium dipicolinic acid (Ca-DPA) release and subsequent rehydration of the core leading to a decrease in the optical density (OD 600nm ) (Setlow 2003, Donnelly et al. 2016). To study the role of CD1065 in germination, we monitored the decrease in the OD 600nm of wild-type and mutant spore suspensions after the addition of TA. When incubated with 1% TA at 37°C for 1 h, we observed a decrease of 50% and 20% in the OD 600nm of the wild-type and mutant spores, respectively (Fig. 3A). The germination rate was partially restored in the complemented strain (Fig. 3A). In addition, when plated in BHI containing 0.1% TA, purified CD1065 mutant spores exhibited a 30-fold defect in spore germination efficiency compared to the wild-type and complemented strains (Fig. 3B). The germination efficiency did not increase after a prolonged incubation (48 h) for both the wild-type and the mutant spores. This result is consistent with the inability of some mutant spores to germinate after 24 h in the presence of 0.1% TA as observed by phase-contrast microscopy (data not shown). These results indicated that CD1065 inactivation likely impairs germination rather than sporulation.

We further monitored the germination process using phase-contrast microscopy. We showed that in the presence of 1% TA, most of the phase-bright spores of the wild-type strain were converted to phase-dark spores after 20 min (Fig. 3C). In contrast, the majority of the mutant spores were still phase-bright after 2 h, suggesting that the CD1065 mutant spores are defective in cortex hydrolysis, the first germination step in C. difficile (Francis et al. 2015). It has been shown that Ca-DPA release occurs immediately after the cortex hydrolysis of the C.

difficile spores (Francis et al. 2015). We tested whether CD1065 mutant spores could release Ca-DPA in response to 1% TA. While the wild-type spores released ~80% of the stored Ca-DPA in response to TA (Fig. S3), only ~20% of the stored Ca-DPA was released under similar conditions for the CD1065 spores. Together, these results suggest that CD1065 mutant from the spore, leading to hydration of the core and to the initiation of outgrowth (Adams et al. 2013, Francis et al. 2015). Our results prompted us to investigate whether the CspBmediated proteolytic activation of SleC is affected in the CD1065 mutant (Adams et al. 2013).

To test this hypothesis, purified spores of the wild-type and the CD1065 mutant strains were treated with 1% TA for 20 min at 37°C. Pro-SleC cleavage was then analysed by Western blotting using an antibody raised against SleC. Interestingly, the amount of Pro-SleC was strongly reduced in the coat of the mutant compared to that of the wild-type strain (Fig. 3D).

After TA treatment, Pro-SleC was cleaved in the spores of the wild-type (Fig. 3D) but not in the CD1065 mutant, even when we increased 3-fold the amount of protein. This result suggests that the CD1065 inactivation interferes with Pro-SleC maturation in response to TA.

Inactivation of CD1065 affects the abundance of major proteins in the spore surface

The lower amount of the pro-SleC protein in the coat extract of the CD1065 mutant spores led us to investigate whether the content of the spore surface layers of this mutant was different from that in the wild-type strain. SDS-PAGE analysis of the spore coat plus exosporium extracts obtained from purified spores showed that several protein bands were absent or present at reduced levels in the mutant spores (Fig. S4A, red arrows). To have a global view of the spore surface proteome of the CD1065 mutant, proteins extracted from the coat/exosporium of spores of the wild-type strain and the mutant were analysed using mass spectrometry. The analysis revealed that the abundance of several spore surface proteins decreased drastically in the CD1065 mutant spores (Table 1). The abundance of the early morphogenetic proteins SpoIVA and SipL (Putnam et al. 2013) and the coat proteins, CotA and CotB (Permpoonpattana et al. 2013), was reduced in the mutant. Among the proteins identified as less abundant in the spore surface layers of the CD1065 mutant when compared to the wild-type strain, we identified two catalases CotD and CotG, the superoxide dismutase SodA, the rubrerythrin CD2845 likely involved in protection against oxidative stress and Table 1). The abundance of other σ K -controlled coat proteins, such as CD3580, CD2400 and CD0597, was also reduced from the coat extract of the CD1065 mutant (Abhyankar et al. 2013). A reduced amount of germination-related proteins, such as the protease CspB and the spore cortex-lytic enzyme SleC, was detected in the mutant, while no differences were observed in the amount of the germinant receptor CspC and the lipoprotein GerS between the wild-type and mutant coat extracts. To validate these results, we performed Western blot experiments using antibodies raised against coat and exosporium proteins. In agreement with the mass spectrometry data, a decreased level of SipL, CotB, CdeC, CdeM, CspB and Pro-SleC was observed in the coat extracts of the mutant spores (Fig. S4B). Both full-length (80 kDa) and cleaved CotE proteins (40 kDa) were detected in the wild-type spores, while the cleaved protein was absent in the coat extract of the mutant spores. Contrary to what we observed by mass spectrometry that we cannot yet explain, a slight decreased level of CspC was detected in the coat extract of the CD1065 mutant spores by Western blot (Fig. S4B). Complementation of the CD1065 mutant restored partially or totally the wild-type levels of these proteins. Together, these results suggest that CD1065 mutation has an effect on the pattern of proteins from the spore surface layers.

CD1065 gene disruption affects the assembly of the surface layers of C. difficile spores

To assess the effect of CD1065 inactivation on the structure of the spore surface layers, the morphology of the CD1065 mutant spores was analysed using transmission electron microscopy (TEM). Previous TEM studies have revealed that the C. difficile spore outermost layers comprise an inner lamellar and an outer electrodense coat layer surrounded by an exosporium (Lawley et al. 2009b, Permpoonpattana et al. 2011b). CD1065 mutant spores analysed by TEM exhibited a major morphological defect. A strong reduction of the surface layers (coat plus exosporium) was observed, although the cortex was maintained (Fig. 4A). On average, the size of the mutant spores (length 617±94 nm/width 476±60 nm) was also smaller than that of the wild-type strain (length 890±324 nm/width 730±74 nm). These data confirmed the first observation performed using phase-contrast microscopy (Fig. 2BC).

The surface layers and the cortex of the mutant spores were 4-fold and 2-fold thinner, respectively, than those of the wild-type spores (Fig. 4B). The production of the phase-grey mutant spores observed using phase-contrast microscopy might be related to the decrease in the thickness of the cortex layer. By contrast, no differences were observed in the size of the spore core (Fig. 4C). Importantly, the complementation of CD1065 restored the assembly of the spore coat and exosporium. In summary, these observations suggested that CD1065 plays an essential role in spore coat assembly and spore morphology.

Disruption of CD1065 increases spore coat permeability

The defective spore coat structure observed in the CD1065 mutant suggested that these spores could have reduced resistance properties. Since the coat layer protects C. difficile spores against lysozyme (Escobar-Cortes et al. 2013), we treated spores of the wild-type, mutant and complemented strains with 250 µg/mL of lysozyme for 1 h. We observed that most of the wild-type spores remained phase-bright after this treatment (Fig. 5A). In contrast, the mutant spores lost the phase-bright property, suggesting that the absence of an intact coat in the CD1065 mutant leads to cortex degradation by lysozyme. Consistent with these results, untreated and lysozyme-treated spores were plated in BHI. Contrary to the wild-type and complemented spores, the CD1065 mutant spores were unable to form colonies on BHI.

However, once in the presence of lysozyme, we restored a level of germination almost similar to that observed in the wild-type or complemented spores (Fig. 5B). These results suggested that lysozyme itself can trigger CD1065 mutant spores germination even in the absence of TA. For the complemented strain, we observed only a partial complementation after lysozyme treatment under the microscope (Fig. 5A). A decreased level of some coat proteins at the spore surface (Fig S4) might slightly affect the coat permeability allowing the lysis of the cortex in at least a fraction of the complemented spores. In addition, by plating the spore suspensions after lysozyme treatment in BHI containing TA, we observed a significant increase of the germination of the CD1065 mutant spores but not in the wild-type and the complemented strains (Fig. 5C). Together, these results suggested that the CD1065 mutant spores are more permeable to lysozyme, which can trigger germination by directly degrading the spore-peptidoglycan cortex.

CD1065 mutation does not block the expression of the σ E -and σ K -target genes

To explain how the CD1065 protein is required for the coat formation, we first tested if CD1065 could be a regulatory protein required for the expression of the genes encoding coat/exosporium proteins. For this purpose, we used qRT-PCR to compare the expression of sporulation-controlled genes in the CD1065 mutant and the wild-type strain after 14 h or 24 h Western blot analyses of coat/exosporium (3) and decoated spore lysate (1 and 2) fractions from the wild-type, CD1065 mutant and complemented strains. The decoated lysate contains the spore core and cortex layers untreated (1) or lysozyme-treated to separate the cortex layer of the core (2). CotB was used as a control for a coat protein.

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of growth in SM. No differences were observed between the two strains in the expression level of different target genes of the sporulation σ factors (Saujet et al. 2013) suggesting that CD1065 is not involved in the regulation of the genes encoding spore coat proteins (Table S4). By using Western blotting, we further analysed the amount of the σ E and σ K -controlled proteins in the crude extracts of sporulating cells harvested after 14 h or 24 h of growth.

Sporulating cells of the CD1065 mutant produced wild-type levels of the σ E -targets (SipL, CotB, CspBA, CspC) and the σ K -targets (SleC, CotE, CdeC and CdeM) (Fig. S4C). These results showed that the deficiencies observed in the coat content of the spores of the CD1065 mutant (Fig. S4B) are not associated with a reduction in the expression of the encoding genes or in protein production. Since this mutant is impaired in its ability to incorporate coat proteins at the spore surface, CD1065 might be a morphogenetic protein required for the presence of key components of the spore coat and exosporium.

CD1065 localizes in the spore surface layers of C. difficile

To determine whether the CD1065 protein is located within the spore envelope, we subjected wild-type and CD1065 mutant spores to a decoating treatment to separate a coat/exosporium soluble fraction and a pellet fraction, which include the core and cortex layers. The pellet was treated with or without lysozyme to hydrolyse the cortex. We performed Western blotting of the different fractions using an antibody raised against the CD1065 protein. This experiment revealed that CD1065 is located only in the coat/exosporium fraction, as CotB used as a coat control (Fig. 6) (Permpoonpattana et al. 2011b). The absence of the signal in the other two fractions indicates that CD1065 is not located in the most internal layers. As CD1065 is found in the coat fraction of mature spores and is rich in lysine, we decided to rename it CotL. Then, we monitored the subcellular localization of this protein during the process of sporulation. We created a C-terminal SNAPtag translational fusion by cloning a fragment encompassing the two promoters and the coding sequence of the cotL gene (P σE/σK(cotL) -cotL-SNAP Cd ) into pFT58 (Pereira et al. 2013, Cassona et al. 2016). The resulting plasmid was introduced in the wild-type strain. Samples of cultures harbouring this translational fusion were collected after 24 h of growth in SM, and the cells were labeled with TMR-Star. Different patterns of CotL localization were detected according to the stages of spore morphogenesis. The fluorescent signal of the P σE/σK(cotL) -cotL-SNAP Cd was detected in the MC compartment during the engulfment and in sporulating cells with phase-dark FS, in 65% and 26% of all the cells in the population with signal, respectively (Fig. 7A). Fluorescent signal around the spore was detected in cells with phase- grey FS (7%) and free spores (2%) (Fig. 7A). No signal was detected in the sporulating cells harbouring only pFT58 (Fig. 7A). To investigate how the inactivation of sigE and sigK influences CotL location, we introduced the P σE/σK(cotL) -cotL-SNAP Cd translational fusion in the respective mutants. As expected, no fluorescent signal was detected in the sigE mutant (Fig. 7A). However, in the sigK mutant, the SNAP fluorescence was detected only in the MC cytoplasm (Fig. 7A), suggesting that the CotL protein cannot be placed and assembled around the FS in the absence of σ K . This is consistent with previous studies showing that the sigK mutant produce spores with no detectable coat layers (Pereira et al. 2013, Pishdadian et al. 2015). Since σ E is active earlier than σ K , we also monitored the location of CotL according to its timing of production. For this, we constructed a second translational fusion containing only the σ K -dependent promoter (P σK(cotL) -cotL-SNAP Cd ). Interestingly, no accumulation of the CotL-SNAP Cd protein was observed in the MC compartment (Fig. 7B). Instead, the fluorescence signal in the sporulating cells was detected first as a cap in the MCP pole during engulfment (6%), and later around the spore in cells with phase-dark and phase-grey FS (58% and 21%, respectively) (Fig. 7B). In free spores (15%), the fluorescent signal was detected around the spore (Fig. 7B). In addition, no fluorescent signal was detected in the sigK mutant but also in the sigE mutant, since σ E is required for sigK expression (Fig. 7B) (Pereira et al. 2013).

Spore coat and exosporium proteins are mis-localized in the cotL mutant cells

To determine the role of CotL in the location of different coat proteins, we constructed C-terminal SNAP-tag translational fusions with known spore coat/exosporium proteins and introduced the resulting plasmids into the wild-type and cotL mutant strains. After 24 h of growth in SM, we determined their patterns of subcellular localization in sporulating cells. As early-synthesized coat proteins belonging to the σ E -regulon, we selected SpoIVA and CotB (Permpoonpattana et al. 2013, Putnam et al. 2013). In the wild-type strain, a signal for the SpoIVA-SNAP Cd signal was detected only in cells with visible phase-dark FS either as a dot, a single cap-like structure in the MCP pole or around the FS (49%, 46% and 5%, respectively) (Fig. 8A). No fluorescent signal was found in free spores (Fig. 8A). The more internal localization of SpoIVA in the spore coat might prevent its detection in mature free spores. In the cotL mutant, the SpoIVA-SNAP Cd protein was found to accumulate either as a dot in the MCP pole of phase-dark FS (88%) or also as dots in the MCP pole and the MC compartment (7%) (Fig. 8A). The position of the red dots overlaps with an apparent accumulation of material visible by phase-contrast microscopy at the MCP pole of the FS. In the cotL mutant, completely free spores were not found. However, contrary to the observed in the wild-type strain, in phase-bright spores of the cotL mutant, a fluorescent signal for the SpoIVA-SNAP Cd fusion was detected in one of the poles (5%) (Fig. 8A, yellow arrow). This signal is consistent with an accumulation of material at the spore pole also visible by phase-contrast microscopy (Fig. 8A). The CotB-SNAP Cd protein fusion was detected in the wild-type strain as a single cap-like structure in the MCP pole (81%), at both FS poles (7%) and around free spores (2%) (Fig S5). In the cotL mutant, CotB-SNAP Cd was mainly found as a dot in the MCP pole of phase-dark FS (97% of the cases). As for the SpoIVA-SNAP Cd fusion, an accumulation of material was detected by phase-contrast microscopy in the same place (Fig. S5). In phasebright spores, this SNAP protein fusion was observed at one pole of the spore (Fig. S5, yellow arrow). These results suggest that cotL inactivation seems to impair the proper assembly around the FS of at least these two σ E -target proteins.

Concerning coat proteins synthesized under σ K control, we selected three proteins, CdeC, CotD and CotE for the construction of translational SNAP fusions (Fig. 8B andS6). The CdeC-SNAP Cd and CotD-SNAP Cd protein fusions were detected in three different patterns in the wild-type strain, i.e., as single cap-like structure in the MCP pole (47% and 50%, respectively), as a double cap in the MCP and mother cell distal (MCD) poles (27% and 17%, respectively), or around the entire FS in sporulating cells with phase-dark FS for CdeC (27%)

and around free spores in the case of CotD (33%) (Fig. 8B andS6A). In the cotL mutant, CdeC-SNAP Cd and CotD-SNAP Cd proteins were detected in 57% and 38% of the sporulating cells as a dot in the MCP pole, respectively (Fig. 8B andS6A). In the other sporulating cells with signal, CdeC-SNAP Cd accumulated in dots in the MCP pole and in the MC compartment (43%). In the cotL mutant free spores, CotD-SNAP Cd fusion was detected at one pole of the spore (Fig. 8B andS6A). For CotE, a non-morphogenetic protein associated with the C.

difficile spore surface (Paredes-Sabja et al. 2014), inactivation of cotL resulted in spots in the MCP pole of the FS (64%) and in MC cytoplasm of cells with either phase-grey or phasebright spores (17% and 19%, respectively), in contrast to the cap-like pattern seen at the MCP in the wild-type strain (Fig. S6B).

Altogether, the results show that the inactivation of cotL gene leads to modifications in the location pattern of both σ E and σ K target proteins suggesting that CotL is required to properly encase the coat proteins around the FS during sporulation.

Discussion

Despite the increased knowledge on the regulatory network controlling sporulation in C. difficile, relatively little is known about the spore composition and the mechanisms that orchestrate the assembly of its surface layers. Except for SpoIVA (Putnam et al. 2013) and SpoVM (Ribis et al. 2017), the key differences in the morphogenetic coat proteins of B.

subtilis and C. difficile make extrapolation difficult. In this study, we propose that the C. difficile coat-associated protein, CotL, is a new morphogenetic coat protein essential for the efficient assembly of the coat and exosporium layers. Spore morphogenetic proteins are key components of the ordered assembly of coat proteins at the spore surface. We observed by TEM analysis that cotL inactivation resulted in a substantial defect in the spore morphogenesis with a strong reduction of the coat and an absence of exosporium layers.

Congruently, the abundance of several spore coat and exosporium proteins is strongly decreased in extracts of the spore surface layers of the cotL mutant compared to those of the wild-type strain (Table 1 and Fig. S4B). However, cotL inactivation did not impair the transcription of sporulation-related genes, or the protein production in sporulating cells. Thus, we infer that CotL plays a morphogenetic rather than a regulatory role in coat formation.

Indeed, we observe that some coat proteins produced under the control of σ E such as SpoIVA and CotB, or of σ K such as CdeC, CotD and CotE are mis-localized in the cotL mutant. In this mutant, they accumulate mainly in the MCP pole or as spots in the MC cytoplasm rather than around the FS. These results suggest that the CotL protein is required to correctly guide and incorporate spore surface proteins around the spore.

The structurally defected coat of the cotL mutant spores has a strong impact on spore function, particularly in their sensitivity to lysozyme and their germination properties. An impaired coat structure strongly affects germination efficiency and spore resistance to lysozyme as observed for the B. subtilis spoVID, safA and cotE mutants (Zheng et al. 1988, Beall et al. 1993, Takamatsu et al. 1999, Costa et al. 2007). Coat-less spores of Clostridium perfringens also do not respond to germinants, but lysozyme can induce germination (Cassier and Ryter 1971). In C. difficile, the morphologic defect of the surface layers of the cotL mutant is probably responsible for an increased spore permeability of lysozyme that can trigger germination by accessing and hydrolysing the spore cortex. The observed germination defect in the cotL mutant is very likely an indirect consequence of a misassembled spore coat and of the reduced amount of CspC, CspB and Pro-SleC proteins in the surface layers of this mutant. The decrease of Csp proteins likely results in a defected maturation of the cortex hydrolase SleC. The inactivation of cotL also leads to the production of a thinner spore cortex Results

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compared to what is seen in the wild-type spores. This is consistent with a slight decrease on heat resistance at 80°C observed for the cotL mutant spores (data not shown).

The expression of the cotL gene is restricted to the MC as expected for a coat protein, and this gene is transcribed under the dual control of the two specific σ factors of the MC, σ E and σ K .

The transcription of cotL is initiated by σ E early during sporulation after asymmetric division, suggesting a potential role of CotL in early FS development, as already observed for SpoIVA and SipL (Putnam et al. 2013). In addition, the transcription of cotL by σ K probably allows the maintenance or increased expression of cotL during the late sporulation stages. Congruently, we observed a reduced abundance of the SNAP protein when produced under the control of the σ E -or σ K -dependent promoter alone compared to its production under the control of both σ E and σ K . In addition, when the cotL-SNAP Cd translational fusion is uniquely transcribed by σ K likely at a reduced level, the CotL-SNAP Cd protein is immediately assembled at the FS surface, while the CotL-SNAP Cd produced under the dual control of σ E and σ K first accumulates in the MC and then assembles around the FS (Fig. 7A andB). The microscopy results also revealed that the CotL-SNAP Cd protein produced under the control of its native promoter is not assembled around the FS in the sigK mutant (Fig. 7A). Consistently with this, the sigK mutant itself is defective in coat polymerization and localization (Pereira et al. 2013, Pishdadian et al. 2015). However, it has been shown that CotB-SNAP Cd (σ E -target) is located at the FS surface in the sigK mutant (Pereira et al. 2013), suggesting that C. difficile coat assembly starts at an early sporulation stage, as in B. subtilis (Henriques and Moran 2007).

Our observations raise the hypothesis that CotL assembly might be dependent on the presence of a σ K -regulated gene product even if we cannot completely exclude that the reduced amount of CotL expressed solely under the control of σ K might be responsible for its direct assembly without accumulation in the MC.

Previously published proteomic studies (Lawley et al. 2009b, Abhyankar et al. 2013, Diaz-Gonzalez et al. 2015) and our proteomic analysis failed to detect the CotL protein in the spore or in the spore surface layers. However, we successfully detected CotL at the spore surface either by fluorescence microscopy using CotL-SNAP Cd fusion or by Western blot analysis of the coat/exosporium extract using an antibody raised against CotL. Since CotL is a lysine-rich protein (24% of the amino acids), this feature might hinder its detection by bottom-up mass spectrometry-based proteomics. Proteins are usually digested into peptides with trypsin, an enzyme that cleaves after lysine and arginine residues and the generated peptides are therefore too small or not suitable with their further identification by mass spectrometry.

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Experimental procedures Bacterial strains and growth conditions

The bacterial strains used in this study are listed in Table S1. Luria-Bertani (LB) medium (1% bacto-tryptone, 0.5% yeast extract, and 0.5% NaCl, pH 7.0) was used for the growth of Escherichia coli. When indicated, ampicillin (100 µg/mL) or chloramphenicol (15 µg/mL) was added to the culture medium. C. difficile strains were grown anaerobically (5% H 2 , 5% CO 2 and 90% N 2 ) at 37°C in brain heart infusion (BHI) medium (Difco). Sporulation assays were performed in sporulation medium (SM) (Wilson et al. 1982). To produce spores, we used an SMC medium containing (per liter): 90 g of Bacto peptone (Bacto TM 211677), 5 g of Proteose peptone (82450, Sigma-Aldrich), 1 g of (NH 4 ) 2 SO 4 and 1.5 g of Tris. When necessary, cefoxitin (25 µg/mL), thiamphenicol (15 µg/mL) and erythromycin (2.5 µg/mL)

were added to the C. difficile cultures. All the primers used in this study are listed in Table S3.

Construction of C. difficile strains

The ClosTron gene knockout system (Heap et al. 2007, Heap et al. 2009) was used to inactivate the gene in 630∆erm. Using the Targetron design software (Sigma-Aldrich), we designed primers to retarget the group II intron of pMTL007-CE5 into the cotL gene in the sense orientation immediately after the 123 rd nucleotide in the coding sequence (Table S3). E. coli HB101(RP4) containing pMTL007-CE5-cotL-123s (pDIA6563) was mated with C. difficile 630∆erm. C. difficile transconjugants were selected by sub-culturing on BHI agar containing thiamphenicol and cefoxitin and plated on BHI agar containing erythromycin.

After extraction of the chromosomal DNA of transconjugants, PCRs were realized to verify the integration of the intron into the cotL gene and the splicing of the group I intron after integration (Fig. S2 and Table S3). This experiment produced strain CDIP782 (630∆erm cotL::erm, Table S1). To complement this mutant, we amplified cotL coding sequence and its promoter region (positions -243 to +438 from the translational start site) using genomic DNA as the template. The PCR product was cloned between the EcoRI and NotI sites of plasmid pMTL84121 (Heap et al. 2009), to produce pDIA6626. This plasmid was transferred by conjugation into C. difficile cotL::erm producing CDIP952 strain (Table S1).

Mapping of the 5 terminus of cotL mRNA

The 5' end of the cotL mRNA was mapped by primer extension using the 5' Rapid Amplification of cDNA Ends (RACE) System (Invitrogen) according to the manufacturer's recommendations. CF127 primer (Table S3) was used to synthesise the first cDNA strand, using the RNAs extracted from 630∆erm after 14 h and 24 h of growth in SM as templates.

The CF131 primer was used for the PCR of the dC-tailed cDNA. The products of the primer extension were analysed by electrophoresis in 1.5% agarose gel. Two different-size products were detected, and TA cloning was used to discriminate the two inserts analysed by sequencing.

RNA extraction and quantitative RT-PCR analysis

To study σ E -or σ F -targets, we harvested the cells from 630∆erm and cotL mutant after 14 h of growth in SM medium, while σ G -and σ K -targets were analysed after 24 h of growth.

RNA was extracted using the FastRNA Pro Blue Kit (MP Biomedicals), according to the manufacturer's instructions. cDNAs synthesis and real-time quantitative PCR were performed as previously described (Saujet et al. 2011, Soutourina et al. 2013). In each sample, the quantity of cDNAs for each gene was normalized to the quantity of cDNAs of the DNApolIII gene. The relative transcript changes were calculated using the 2 -2∆∆Ct method as previously described (Livak and Schmittgen 2001). The primers used for each marker are listed in Table S3.

Transcriptional and translational SNAP Cd fusions

To construct transcriptional fusions, pFT47 containing the SNAP Cd sequence was used (Pereira et al. 2013, Cassona et al. 2016). The promoter region of the cotL gene (P1 plus P2) was PCR amplified using genomic DNA and primer pairs CF120/CF121 to produce a 250 bp PCR product (position -243 to +19 from the TSS of P2). To construct the fusion bearing only P2, the σ K -dependent promoter of cotL, a 90 bp region was amplified using the CF133/CF121

primer pairs (position -75 to +19 from the TSS of P2). The DNA fragments were inserted between the EcoRI and XhoI sites of pFT47 to obtain pDIA6561 and pDIA6613 (Table S1).

To construct the fusion bearing only promoter P1, we removed promoter P2 by inverse PCR using pDIA6561 as a template and the CF206/CF207 primer pair. To create a C-terminal SNAP-tag translational fusion, the promoter (P1 plus P2 or P2 alone) and coding regions of (CdeC) were amplified from genomic DNA using adapted primer pairs (Table S3).

CF124/CF202 and CF203/CF204 pairs were used to amplify the promoter present upstream of CD2399 the first gene of the cotD operon and the coding region of CD2401 (cotD), respectively. The PCR products were inserted into pFT58 using Gibson Assembly. The plasmids obtained were introduced in E. coli HB101 (RP4) and transferred by conjugation to the C. difficile 630∆erm and cotL::erm mutant strains (Table S1).

Fluorescence microscopy and image analysis

For SNAP labelling, the TMR-Star substrate (New England Biolabs) was added to the cells at a final concentration of 250 nM, and the mixture was incubated for 30 min in the dark and in anaerobiosis. After labelling, the cells were collected by centrifugation (4000xg for 3 min at room temperature), washed 3 times with 1 mL of PBS, and finally resuspended in 1 mL of PBS containing or lacking the membrane dye Mitotracker Green (MTG; 0.5 µg/mL) (Molecular Probes, Invitrogen). For phase-contrast and fluorescence microscopy, spores or cells were mounted on 1.7% agarose-coated glass slides to keep the cells immobilized and to obtain sharper images (Glaser et al., 1997). The images were taken with exposure times of 200 ms for MTG and 600 ms for SNAP. Concerning the transcriptional fusions, cells were observed using a Nikon Eclipse TI-E microscope 60x Objective and captured with a CoolSNAP HQ2 Camera. For the translational fusions, cells were observed using a Delta Vision Elite microscope equipped with an Olympus 100x objective and captured with sCMOS camera. The images were analysed using ImageJ.

Spore production and sporulation tests

SMC liquid medium inoculated with C. difficile was incubated overnight at 37°C.

Two hundred microlitres from this culture plated in SMC agar were incubated at 37°C for 7 days. Spores scraped off with water were incubated for 7 days at 4°C. Cell fragments and spores were separated by centrifugation using a HistoDenz (Sigma-Aldrich) gradient (Dembek et al. 2013). Suspensions (>99% of spores) are free of vegetative cells, sporulating cells and cell debris as determined by phase-contrast microscopy. The spore suspension was quantified by measuring the OD 600nm.

Overnight cultures grown at 37ºC in BHI were used to inoculate SM media at a dilution of Results 97 1:200. At specific time points, 1 mL of culture was withdrawn, serially diluted in BHI and plated before and after heat treatment (30 min, at 60ºC), in order to determine the total and heat-resistant colony forming units (CFU). Samples were plated onto BHI supplemented with 0.1% TA (Sigma-Aldrich) to promote efficient spore germination (Wilson et al. 1982).

Germination assay

Approximately 10 8 spores (equivalent of 0.4 OD 600nm units) were resuspended in 200 µL of water. The spore suspension was heat-treated at 65ºC during 10 min, and serial dilutions were plated on BHI containing 0.1% TA to determine the number of spores that are able to germinate. The CFU were monitored at 24 h and 48 h. At least three biological replicates were performed using at least two independent spore preparations.

The germination of the spores was also monitored by OD 600nm as previously described (Donnelly et al. 2016) and by phase-contrast microscopy as described previously (Kong et al. 2011). For optical measurement, spores resuspended in BHI were exposed to 1% TA, and the OD 600nm was measured every 5 min for 1 h. For the phase-contrast microscopy, purified spores were resuspended to an OD 600nm of 0.4 in BHI containing 1% TA and incubated under anaerobic conditions for 2 h at 37°C. At each time point, samples were pelleted by centrifugation, washed with PBS and fixed with 2% paraformaldehyde for 15 min at room temperature. Fixed samples were washed with PBS once more and dried onto glass slides.

Phase-contrast microscopy was conducted using an Axioskop microscope (Zeiss) with a 100x oil immersion lens. The images were captured using an Axiocam 105 colour (Zeiss).

Ca-DPA release and SleC cleavage assays and lysozyme resistance tests

Measurement of the DPA release was performed as previously described (Francis et al. 2013a) from purified spores incubated at 37°C in PBS supplemented with 1% TA and 10 mM glycine for 1 h. Equal aliquots were incubated at 100°C as a measure of 100% DPA release (positive control) or incubated at 37°C in PBS without the germinants (negative control). SleC cleavage assay was performed as previously described (Kevorkian et al. 2016) from spores suspended for 20 min in BHI containing 1% TA. Lysozyme resistance of C.

difficile spores was measured by suspending 10 8 spores in 200 µL of PBS in the presence or absence of 250 µg/mL of lysozyme. Spores were incubated for 1 h at 37°C with shaking (300 rpm), and the samples were visualized by phase-contrast microscopy and plated onto BHI agar containing 0.1% TA and incubated anaerobically for 24 h at 37°C.

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Transmission electron microscopy

To analyse the ultrastructure by TEM, purified spores of the wild-type, cotL mutant and complemented strains were fixed with 2.5% of glutaraldehyde (Sigma Aldrich) in 0.1 M HEPES, pH 7.2. Post fixation was conducted with 1% osmium tetroxide (Merck) and 1.5% ferro-cyanide (Sigma Aldrich) in 0.1 M HEPES. After dehydration by a graded series of ethanol, spores were infiltrated with epoxy resin. Seventy-nanometer sections were collected on formvar coated slot grids (EMS) and were contrasted with 4% uranyl acetate and Reynolds lead citrate. The stained sections were observed using a Tecnai spirit FEI operated at 120 kV.

The images were acquired using an FEI Eagle digital camera.

Coat extraction and Western blot analysis

To analyse the profile of the coat and exosporium proteins, C. difficile spores were resuspended in extraction buffer (0.125 mM Tris-HCl, 5% β-mercaptoethanol, 2% SDS, 0.5 mM dithiothreitol (DTT) and 5% glycerol, pH 6.8) containing 0.025% bromophenol blue and boiled as previously described (Costa et al. 2006). For coat and cortex fractionation, the spores were first resuspended in extraction buffer, boiled and centrifuged to recover the supernatant corresponding to the coat fraction (including the coat and exosporium layers).

Pellets were treated with or without 2 mg/mL of lysozyme for 2 h at 37°C. For sporulating cells, whole cell lysates were prepared as described previously (Fagan and Fairweather 2011).

Briefly, sporulating cultures of C. difficile were harvested by centrifugation at 5000 rpm for 10 min at 4°C, and the pellets were frozen at -20°C. The bacteria were thawed and resuspended in PBS containing 0.12 µg/mL of DNase I to an OD 600nm of 40. An equal volume of 2X extraction buffer containing 0.025% bromophenol blue was added to the protein samples (Laemmli 1970), which were resolved by SDS-PAGE and visualized by Coomassie brilliant blue R-250 staining (Bio-Rad). Proteins were transferred to a PVDF membrane (0.45 µm) and subjected to immunoblot analyses as described previously (Serrano et al. 1999). antibodies conjugated to horseradish peroxidase (Sigma) were used at a 1:5000 dilution.

Mouse polyclonal antibody raised against

Immunoblots were developed with enhanced chemiluminescence reagents (Thermo Scientific).

Antibody production

To construct cotL expression the plasmid, the coding sequencing of the cotL gene was PCR amplified using genomic DNA and primer pairs CF184/CF185 to produce a 500 bp PCR product. The DNA fragment was cloned between the NdeI and XhoI sites of pET20 to obtain pDIA6636 (Table S1). After verification by sequencing, the plasmid was introduced in E.coli BL21 (DE3) yielding ECO832 (Table S1).

Mass spectrometry of the spore coat and exosporium layers

C. difficile spores were resuspended in extraction buffer and boiled as previously described (Costa et al. 2006). Coat and exosporium proteins were recovered by centrifugation and precipitated with 10% TCA. For spore protein digestion, protein pellets were resuspended in urea 8 M and NH 4 HCO 3 100 mM and sonicated on ice. Subsequently, 50 µg of total proteins were reduced with TCEP 10 mM (646547 -Sigma) for 30 min and then alkylated with iodoacetamide 20 mM (I114 -Sigma) for 1 h. Proteins were digested with rLys-C 1 µg (V1671, Promega) for 4 h at 37°C and then digested with trypsin 1 µg (V5111, Promega) overnight at 37°C. The digestion was terminated with formic acid 4%, and the peptides were desalted using a C18 Sep-Pak Cartridge (WAT054955, Waters). The peptides were eluted successively with 50% Acetonitrile (ACN)/0.1% Formic acid (FA) and 80% ACN/0.1% FA before being dried in a vacuum centrifuge. The peptides were resuspended in 2% ACN /0.1% FA.

For LC-MS/MS analysis, a nanochromatographic system (Proxeon EASY-nLC 1000, Thermo Fisher Scientific) was coupled online to a Q Exactive TM HF Mass Spectrometer (Thermo Fisher Scientific). For each sample, 1 µg of peptides was injected onto a 50-cm homemade C18 column (1.9 µm particles, 100 Å pore size, ReproSil-Pur Basic C18, Dr.

Maisch GmbH) and separated with a multi-step gradient from 2 to 45% ACN in 105 min at a Results 100 flow rate of 250 nL/min over 132 min. The column temperature was set to 60°C. The data were acquired as previously described (Lago et al. 2017).

Raw data were analysed using MaxQuant software version 1.5.5.1 (Cox and Mann 2008) and the Andromeda search engine (Cox et al. 2011). The MS/MS spectra were searched against an internal C. difficile database containing 3 957 proteins, and the contaminant file included in MaxQuant. The database search parameters were fixed as described (Lago et al. 2017). For the quantification, skip normalization option was used. For the statistical analysis of one biological condition versus the other, it was used the same data analysis pipeline than in (Jeannin et al. 2018). Exceptionally, the proteins exhibiting lower than 2 LFQ value in both conditions were discarded from the list to avoid misidentified proteins (Jeannin et al. 2018).

The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD010359 (Vizcaino et al. 2016).

II. Complementary results on Spore coat morphogenesis II.1. Investigation of potencial CotL partners

Our study revealed that CotL is an important morphogenetic factor of the C. difficile spore. In fact, CotL is located in the spore surface layers coat/exosporium and is required for the proper localization of several coat proteins around the FS during spore development. We showed that CotL is not apparently involved in the regulation of genes encoding coat proteins, since a cotL mutant is able to produce wild-type levels of any of the coat proteins tested.

Altogether, our results showed that CotL protein is crucial for the assembly of the spore coat, playing a morphogenetic rather than a regulatory role in the spore formation.

Spore coat morphogenetic proteins of B. subtilis are typically known to interact with and recruit other coat components (Henriques andMoran 2007, McKenney et al. 2013). SpoIVA localization to the spore occurs through a direct interaction with SpoVM (Price and Losick 1999). Moreover, SpoIVA interacts with SpoVID, and a physical association between this last protein and the morphogenetic proteins of the inner and outer coat layers, SafA and CotE, also occurs [START_REF] Ramamurthi | Peptide anchoring spore coat assembly to the outer forespore membrane in Bacillus subtilis[END_REF], Mullerova et al. 2009).

The requirement of C. difficile CotL for proper localization of several coat proteins at the spore surface suggests that this protein might interact and recruit these coat components.

In

II.1.a. Identification of proteins interacting with CotL by BACTH

The inactivation of cotL leads to an almost uncoated spore and SpoIVA and SipL are the first proteins starting the assembly of the coat layer. Therefore, we decided to test first whether CotL could interact with these two proteins. Then, we investigated possible interactions between CotL and other coat components such as CotB and CotA, whose expression is controlled by σ E and σ K , respectively and present mainly in the coat layer; CotE, controlled by σ K , which is an enzymatic component of the spore exosporium; and lastly CdeC, a member of the σ K regulon required as a morphogenetic protein of the exosporium.

His-CotL

We investigated possible interactions between CotL and these C. difficile spore proteins by using a bacterial two hybrid system (BACTH). This system is based on the reconstitution of an adenylate-cyclase from the N-terminal (T25 fragment) and C-terminal (T18 fragment) parts when the two proteins fused to these parts interact to each other (Karimova et al. 1998).

The production of cAMP by the reconstituted enzyme leads to the expression of lacZ allowing detection of the interaction. CotL, SipL, CotB, CotA, CotE and CdeC were fused to T18 and T25 both in N-and C-terminal positions. Unfortunatelly, we were unable to construct the plasmids to test the interaction between CotL and SpoIVA proteins. The interaction of fusion partners was investigated by testing β-galactosidase activity. Negative control experiments were carried out using the empty plasmids pKNT25 and pUT18 (Table 1). According to the quantification of the different combinations tested, β-galactosidase activity resulting from an interaction between CotL and SipL was detected in five different combinations out of eight (Table 1). A possible interaction between CotL and CotB, and between CotL and CotE has been detected only in one combination, and in both of them, the C-terminal part of CotL is fused to the T25 part. No β-galactosidase activity was detected in any combination of the CotL-CotA and CotL-CdeC. A CotL self-interaction was also investigated but no βgalactosidase activity was detected. These results suggest that CotL might interact with SipL and CotB and CotE might be possible partners of this protein to a lesser extent. Finally, no interaction was observed within CotL and between CotL and CotA or CdeC.

II.1.b. Coexpression of spore coat proteins and pull down assay in E. coli

In order to confirm the positive interactions obtained using the BACTH system, we decided to use another biochemical assay. We used for this task the pETDuet system that allow to coexpress genes encoding two proteins, that make possible interaction inside the cells and consequently purification of the protein complex. In this case, one protein carries the polyhistidine-tag and could be purified by affinity on a Cobalt (Co 2+ ) column and the second is produced as a native protein and can be pulled down. We used this assay to search for interactions between CotL and SipL, CotB or CotE. We also analysed a possible interaction between CotL and SpoIVA. As positive control, we coexpressed spoIVA and sipL since these proteins have been previoulsy shown to interact (Putnam et al. 2013). Immunoblot analysis of E. coli whole cells lysates using antibodies raised against the His-tag or specific coat protein confirmed that all proteins were produced in E. coli (Figure 32 and 33). To identify potential 32A and 33A). By contrast, His-tagged CotL proteins were always detected in the elution fractions, showing a specific binding onto the affinity column. The obtained results showed that, in our conditions, SpoIVA was pulled down with His-tagged SipL after coexpression of both genes in E. coli (Figure 32C). However, CotL was not pulled down when co-produced with His-SipL (Figure 32A and B), His-CotB (Figure 33A and B) or His-CotE (Figure 33A and C). In addition, SpoIVA was not pulled down with His-CotL (Figure 32A andC). Thus, even if a positive interaction was detected by BACTH assay, we were not able to confirm this result using pull down assays.

II.2. The CD2399 gene

II.2.a. CD2399 and CD0596 are σ K -controlled genes

The expression of most of the genes encoding proteins required for coat synthesis and assembly in C. difficile are transcribed by σ E and σ K and regulated by the DNA binding protein SpoIIID (Fimlaid et al. 2013, Saujet et al. 2013). In order to identify additional genes whose products might play a role in spore coat, we selected another gene, CD2399, which belongs to the σ K regulon (Saujet et al. 2013) and 53% identity to CotJB and CotJC of B. subtilis, respectively. CD0597 protein is removed from the spore surface by sonication and the inactivation of its encoding gene does not affect the ultrastructure of the spore suggesting that this spore protein is located in the exosporium layer (Permpoonpattana et al. 2013, Diaz-Gonzalez et al. 2015). We noted that CotCB (CD2400) and CotD (CD0598) share 71% of identity.

Previous transcriptome studies showed that the expression of CD2399, cotJB2 and cotD in one hand and CD0596, cotJB1 and cotCB in other hand is 100-and 50-fold downregulated in a sigK mutant compared to the wild-type strain, respectively (Saujet et al. 2013).

These results combined with the presence of a sequence similar to σ K consensus elements upstream of the transcriptional start site of both CD2399 and CD0596 genes strongly suggest that their expression is controlled by σ K (Figure 34) (Saujet et al. 2013). Since no promoter was identified upstream of cotJB2-cotD genes and of cotJB1-cotCB, CD2399-coJB2-cotD and CD0596-coJB1-cotCB might form two tricistronic operons.

II.2.b. CD2399 is not required for sporulation

In order to test the role of CD2399 in the spore formation, the encoding gene CD2399

was inactivated using the allele couple exchange (ACE) mutagenesis system (Cartman et al. 2012). When we analyzed the morphology of the strain by phase contrast microscopy after 24 h of growth in sporulation medium (SM), we observed a similar percentage of sporulating cells between the CD2399 mutant and wild-type strains (Figure 35A). The efficiency of heatresistant spore formation was also tested and the wild-type and the CD2399 mutant produced 1.37x10 7 and 1.67x10 7 heat-resistant spores/ml, respectively (Figure 35B). These data suggest that the CD2399 mutant sporulates similarly than the wild-type strain.

II.2.c. CD2399 mutant spores have a germination delay

We also evaluated whether inactivation of CD2399 affected spore germination. We monitored the decrease in the OD 600nm of the wild-type and mutant spore suspensions after the addition of 1% taurocholate (TA). After 5 min in the presence of TA, we observed a decrease of 40% in the OD 600nm of the wild-type spores, while the decrease is of 25% for the CD2399 mutant suggesting a delay in the germination rate (Figure 36A). However, the colony formation efficiency of CD2399 mutant spores in BHI agar plates with TA was similar to that of the wild-type strain (Figure 36B). We further monitored the germination process using phase contrast microscopy. In the wild-type strain, only phase dark spores were observed after

Length Width

Results 109 20 min in the presence of the germinant. However, in the same conditions, only 75% of the CD2399 mutant spores were converted into phase dark spores (Figure 36C). Moreover, four hours after germination induction, outgrowth was observed in both strains but few phase bright spores were still detected in the CD2399 mutant. Altogether these results suggest that part of the CD2399 mutant spores have a germination delay, whereas they are able to germinate since no differences are observed when the same amount of wild-type and CD2399 mutant spores are platted in BHI supplemented with TA.

II.2.d. The inactivation of CD2399 does not affect the ultrastructure and resistance of the spore

Because σ K plays an important role in the assembly of the spore coat and CD2399

gene is a member of the σ K regulon, we compared the ultrastructure and the polypeptide composition of the spore surface layers of the wild-type and the CD2399 mutant. Spores from the wild-type and CD2399 mutant were first purified and analysed by transmission electron microscopy (TEM). TEM analysis showed that CD2399 mutant spores have the same size and a structure similar to the wild-type; moreover no obvious differences were observed in the spore coat layers, in the cortex and in the core (Figure 37). Spore surface proteins were then extracted and we found no significative difference of the protein profiles between the CD2399 mutant and the wild-type (Figure 38A). We further analyzed by Western blot the amount of some spore coat and exosporium proteins, such as the cortex hydrolase SleC, the peroxiredoxin-chitinase CotE and the exosporium morphogenetic protein CdeM, and we observed a slight decrease of the level of these proteins in the ∆CD2399 mutant when compared to the wild-type strain (Figure 38B). Altogether, these observations suggest that the deletion of CD2399 does not affect the assembly of the spore surface layers, even if the amount of the tested coat/exosporium proteins slightly decreased in the mutant. When we tested the resistance of the wild-type and the CD2399 mutant spores to lysozyme, ethanol and to a higher temperature (80 °C), we observed that CD2399 mutant spores are as resistant as the wild-type spores to these conditions tested (Figure 39).

Evidences suggest that all products of the operon to which CD2399 belongs are components of the C. difficile spore coat. In fact in B. subtilis, the members of the similar cotJ operon (cotJA and cotJC) encode spore coat proteins located at the undercoat/basement layer.

However the exact localization of CD2399 in the coat layers of C. difficile is not yet clear.

Moreover the differences in the genetic control of the two operons, i.e., controlled by σ E and involved in earlier stages of coat assembly. Our initial data indicated that CD2399 is not required for spore formation. The CD2399 mutant spores are structurally similar and present the same resistance properties to lysozyme, heat and ethanol than that of the wild-type strain. The only differences observed in the CD2399 mutant when compared to the wild-type strain, concerned both the small delay in germination and the slight decrease in the level of some coat proteins. In B. subtilis, the inactivation of any of the members from cotJ operon results in the formation of spores with a normal morphology and no detectable resistance defiency [START_REF] Henriques | Characterization of cotJ, a sigma E-controlled operon affecting the polypeptide composition of the coat of Bacillus subtilis spores[END_REF][START_REF] Seyler | Assembly and interactions of cotJ-encoded proteins, constituents of the inner layers of the Bacillus subtilis spore coat[END_REF]). However, in B. subtilis, CotJA and CotJC proteins are not detected in the coat of the cotJC and the cotJA mutants respectively, suggesting that CotJA might interact with CotJC for their assembly into the spore coat [START_REF] Seyler | Assembly and interactions of cotJ-encoded proteins, constituents of the inner layers of the Bacillus subtilis spore coat[END_REF]. In fact, an interaction between CotJA and CotJC has been observed [START_REF] Seyler | Assembly and interactions of cotJ-encoded proteins, constituents of the inner layers of the Bacillus subtilis spore coat[END_REF]

CFUs/mL

Wild-type ΔCD2399

O 2 tolerance and ROS protection in C. difficile

After spore germination, C. difficile vegetative cells multiply and colonize the intestine. Then, the cells produce virulence factors including the two toxins, TcdA and TcdB, which cause the pathology associated with CDI. These toxins are responsible for alterations in the actin cytoskeleton of the intestinal epithelial cells, inducing intestinal-cell lysis and triggering an important inflammation (Abt et al. 2016a, Janoir 2016). The inflammation process results in the production of ROS, NO and RNS at bactericidal concentrations by the host immune system (Britton andYoung 2014, Theriot andYoung 2015). Apart from the oxidative stress, O 2 is also an important stress that C. difficile might face during gut colonization. Indeed, although a healthy gastrointestinal tract with a diverse microbiota is regarded as mainly anoxic, an increasing O 2 gradient from the lumen of the colon toward the intestinal epithelium is present [START_REF] Marteyn | Breathing life into pathogens: the influence of oxygen on bacterial virulence and host responses in the gastrointestinal tract[END_REF], Kelly et al. 2015a). Additionally, antibiotic-induced disruption of the host microbiota leads to an increase in O 2 within the gut (Kelly et al. 2015a[START_REF] Rivera-Chavez | Depletion of Butyrate-Producing Clostridia from the Gut Microbiota Drives an Aerobic Luminal Expansion of Salmonella[END_REF]). Thus, O 2 concentration fluctuations in the gastrointestinal tract presents a challenge to anaerobic bacteria such as C. difficile [START_REF] Marteyn | Breathing life into pathogens: the influence of oxygen on bacterial virulence and host responses in the gastrointestinal tract[END_REF]. Recently it was demonstrated that C. difficile is capable of growth in non-strict anaerobic conditions (1-3% O 2 ) (Kint et al. 2017, Giordano et al. 2018a) indicating this bacterium encodes an arsenal of proteins involved in O 2 detoxification and protection.

However, we still know little about the proteins involved in the ability of C. difficile to tolerate O 2 . In other organisms, both flavodiiron proteins (FDPs) and rubrerythrines (Rbrs) have been shown to play an important role protecting the cells from O 2 , oxidative or nitrosative stress, suggesting that those proteins might play a role in these responses in C. difficile [START_REF] Sztukowska | Role of rubrerythrin in the oxidative stress response of Porphyromonas gingivalis[END_REF][START_REF] Wildschut | Rubredoxin:oxygen oxidoreductase enhances survival of Desulfovibrio vulgaris hildenborough under microaerophilic conditions[END_REF], Hillmann et al. 2009b, Riebe et al. 2009). FDPs are enzymes composed of a minimal core containing both a non-heme diiron catalytic center and a flavodoxin FMN-containing domain located at the N-terminal and Cterminal part of the protein, respectively [START_REF] Romao | The dual function of flavodiiron proteins: oxygen and/or nitric oxide reductases[END_REF], Folgosa et al. 2018a). These

112

The main activity for Rbrs and revRbrs is as peroxidase, linked to NAD(P)H oxidation, but O 2 -reductase activity has been also reported [START_REF] Coulter | NADH peroxidase activity of rubrerythrin[END_REF], Weinberg et al. 2004, Kurtz 2006). To shuttle electrons, both FDPs and revRbrs require other proteins like a rubredoxin (Rd) and NADH/FAD dependent oxidoreductases [START_REF] Kawasaki | O2 and reactive oxygen species detoxification complex, composed of O2-responsive NADH:rubredoxin oxidoreductase-flavoprotein A2-desulfoferrodoxin operon enzymes, rubperoxin, and rubredoxin, in Clostridium acetobutylicum[END_REF], Riebe et al. 2009).

Interestingly, the genes encoding both revRbrs and FDPs are positively controlled by σ B (Kint et al. 2017). This sigma factor is crucial for the management of oxidative and nitrosative stresses and to O 2 tolerance (Kint et al. 2017). From all enzymes protecting the cells from O 2 , oxidative or nitrosative stress, only the FDP protein, CD1623, has been functionally characterised as an O 2 -reductase (Folgosa et al. 2018b).

In the present study, we sought to functionally characterize the second FDP, CD1157, and the two revRbrs of C. difficile and to determine the physiological role of these proteins in O 2 tolerance and oxidative stress resistance of this anaerobic pathogen. The expression of CD1474 has been showed to be only σ B -dependent (Kint et al, in Annex). To analyse the expression of CD1524 and CD1623, the promoter region of each gene was fused to the gene encoding a fluorescent reporter SNAP Cd into the vector pFT47 (Pereira et al. 2013, Cassona et al. 2016). The resulting plasmids were transferred into the wild-type strain and the sigB mutant. After 16 h of growth in BHI, cells were labelled with TMR-Star.

Following labelling, we observed an expression of the P (CD1524) -and the P (CD1623) -SNAP Cd fusions in the wild-type strain in 82% and 58% of the cells, respectively (Figure 40C andD).

By contrast, no fluorescent signal was detected in the cells of the sigB mutant.

The presence of two promoters upstream of the CD1157 gene suggests that its expression might be controlled by both σ A and σ B . To monitor the expression of CD1157, we engineered a transcriptional fusion between the entire promoter region containing both promoters and the SNAP Cd gene (Pereira et al. 2013, Cassona et al. 2016) and transferred the resulting plasmid into the wild-type strain and the sigB mutant. We observed an expression of the P σ A σ B (CD1157) -SNAP Cd fusion in 12% of the cells in the wild-type strain after 16 h of growth in BHI (Figure 41B), while in the sigB mutant, the expression of P σ A σ B (CD1157) -SNAP Cd is observed, but to a lower extent (2% of cells) (Figure 41B). To confirm the dual control of CD1157 expression, we fused the promoter region harboring either the σ A consensus sequence P σ A (CD1157) or the σ B consensus sequence P σ B (CD1157) to the SNAP Cd encoding gene. The expression of P σ A (CD1157) -SNAP Cd was observed in both the wild-type and the sigB mutant strains in 12% and 2% of the cells, respectively (Figure 41C). By contrast, no fluorescence was detected for P σ B (CD1157) -

114

SNAP Cd in the sigB mutant (Figure 41D) whereas the expression from this fusion was observed in the 7% of the cells of the wild-type strain. Altogether, these results indicate that CD1157 expression is related to both the σ A -and σ B -dependent promoters while CD1524 and CD1623 expression is only σ B -dependent as already observed for CD1474 encoding the other revRbr (Kint et al, in Annex).

I.2. FDP and revRbrs sequence analysis

The CD1157 gene encodes a Class A FDP (see Introduction, chapter 3, section: 1.2.b Flavodiiron proteins) with 397 amino acid residues harbouring the minimal core that is common to all FDPs so far analysed (Folgosa et al. 2018a). The N-terminal part shares similarities with the fold of metallo-ß-lactamase and contains a non-heme diiron center, which is the catalytic site involved in the reduction of O 2 or NO. The C-terminal part is composed of a flavodoxin FMN-containing domain ensuring the electron transfer (Figure 42). Amino acids involved in the coordination of the diiron center in FDPs, including histidines (His 83 , His 88 , His 149 , His 229 ) aspartates (Asp 87 , Asp 168 ) and glutamates (Glu 85 ) are conserved in CD1157. As other FDPs, CD1157 likely receives electrons from a NAD(P)H-OR to reduce O 2 and/or NO.

The two revRbrs, CD1474 and CD1524, share 96% of amino acid identity (Figure 43). Two domains, found in inverted position in classical Rbrs, compose these proteins: an electron transfer Rd domain binding a Fe(Cys) 4 center at the N-terminal and a diiron catalytic center at the C-terminal part [START_REF] Demare | The structure of Desulfovibrio vulgaris rubrerythrin reveals a unique combination of rubredoxin-like FeS4 and ferritin-like diiron domains[END_REF]. The Rd domain of 33 amino acids contains two CysXXCys motifs (Cys6/Cys9, Cys22/Cys25) separated by 12 residues, which is typical of short-spaced Rd domain found in Rbrs [START_REF] Cardenas | Aerobic Lineage of the Oxidative Stress Response Protein Rubrerythrin Emerged in an Ancient Microaerobic, (Hyper)Thermophilic Environment[END_REF].

I.3. Protein characterization and redox properties

The FDP CD1157, and the two revRbrs, CD1474 and CD1524, were produced as nontagged proteins in E. coli grown in M9 medium supplemented with iron in the presence of IPTG. The proteins were then purified through several chromatographic steps. The molecular masses determined from the SDS-PAGE, ~ 45 kDa for CD1157 and ~ 22 kDa for the two revRbrs, were in agreement with those calculated from the respective amino acids sequence, 44 and 20 kDa (Figure 44). Gel filtration of the purified recombinant proteins (data not shown) revealed that CD1157 behaves as a homodimer in solution, presenting a molecular mass of ~ 74.7 kDa while both revRbrs are isolated as tetramers in solution with determined molecular masses of ~ 79.2 kDa for the CD1474 and ~ 79.8 kDa for the CD1524. FDPs so far for efficient electron transfer between the FMN of one monomer and the diiron center of the other (Folgosa et al. 2018a). The same configuration, homodimers or homotetramers is usually found in Rbrs [START_REF] Tempel | Structural genomics of Pyrococcus furiosus: X-ray crystallography reveals 3D domain swapping in rubrerythrin[END_REF][START_REF] Pinto | Desulforubrerythrin from Campylobacter jejuni, a novel multidomain protein[END_REF]. Reverse phase HPLC analysis identified FMN as the flavin cofactor of CD1157 (data not shown), which is usually observed in the flavodoxin core domain of FDPs [START_REF] Romao | The dual function of flavodiiron proteins: oxygen and/or nitric oxide reductases[END_REF], Folgosa et al. 2018a). In addition, the UV-visible absorbance spectrum of CD1157 is characteristic of a protein containing flavin as cofactor, exhibiting maxima at 380 nm and 450 nm (Figure 45A). By contrast, the UVvisible absorbance spectra of CD1474 and CD1524 are characteristic of proteins containing a Rd center, exhibiting maxima at 380 nm and 490 nm (Figure 45B andC). For all proteins analyzed, the presence of a diiron site is not detected due to its low molar absorptivity [START_REF] Solomon | Geometric and electronic structure/function correlations in non-heme iron enzymes[END_REF].

Finally, analyses of CD1157 showed a ratio for iron/FMN/protein monomer of 1/0.3/1. These values are lower than expected (2/1/1) suggesting an uncomplete incorporation of iron and FMN in the protein even after FMN supplementation following purification.

CD1474 and CD1524 showed an iron/protein monomer ratio of 1.3/1 and 2.8/1.1, respectively. CD1524 appeared to be fully loaded with iron while the ratio was lower than expected for CD1474.

The reduction potentials of the Rd center for the revRbrs CD1474 and CD1524 were obtained by redox titrations monitored by visible spectroscopy (Figure 46) in an anaerobic chamber at pH 7.5. The Rd site was monitored at 490 and 556 nm and a potential of -10 mV and +10 mV were obtained respectively for CD1524 and CD1474. Redox titration of the flavin center of CD1157 will be done in future experiments.

I.4. CD1474 and CD1524 reduce both H 2 O 2 and O 2 while CD1157 reduces only O 2

Having established the presence of the FMN and iron cofactors and determined the redox properties of each protein, we addressed the catalytic activities of CD1157 and of the two revRbrs. The O 2 -and H 2 O 2 -reductase activities of the three enzymes were determined using NADH as the primary electron donor. Purified CD1157, CD1474 and CD1524 (at 30 µM) alone did not show detectable NADH oxidase activity (data not shown). Since the C. difficile physiological electron donors of these proteins are still unknown, external partners were used to perform the assays. We found that a NROR and a Rd from E. coli were able to

I.5. The function of the revRbrs and the FDPs, in the management of O 2 tolerance

To study in C. difficile the physiological role of the two revRbrs and the two FDPs, including the CD1623, already characterized as an O 2 -reductase (Folgosa et al. 2018b), we generated the corresponding mutants. We inactivated the CD1157 gene using the Clostron system, which enabled the insertion of a type I intron after the 225 th nucleotide in sense orientation (Figure S1). Using the allele chromosomic exchange (ACE), we deleted the CD1474, CD1524 and CD1623 open reading frames yielding ∆CD1474, ∆CD1524 and ∆CD1623 single mutants. We also generated ∆CD1474-∆CD1524 and ∆CD1623-CD1157 double mutants (Figure S1). As the four proteins under study possess an O 2 -reductase activity,

we assessed the ability of the different mutants to grow when exposed to low O 2 tension.

Concerning the revRbrs, no difference in growth between the wild-type and the double ∆CD1474-∆CD1524 mutant strains was observed in anaerobiosis or when exposed to 0.1 % of O 2 (Figure 49). By contrast, a growth defect was observed for the ∆CD1474-∆CD1524

double mutant in the presence of 0.2 % of O 2 (Figure 49). The growth of this mutant was even more affected when exposed to 0.4 % of O 2, when compared to the wild-type. Of note, we did not observe any growth defect for the ∆CD1474 or ∆CD1524 single mutant whatever the O 2 tension used suggesting a functional redundancy between these two proteins. Accordingly, wild-type gene was enough to fully restore the growth in presence of low O 2 tension (Figure 49). For the FDPs, a growth defect was observed for the CD1157 mutant in the presence of 0.4 % of O 2 (Figure 50). In the same conditions, even if no major growth defect was observed for the ∆CD1623 mutant, cells seem to be affected since they did not grow in the middle of the spot for some dilutions. To assess if O 2 -reductase activity of CD1157 and CD1623 could be additive, the ∆CD1623-CD1157 double mutant was exposed to 0.4% O 2 . The growth of the double mutant was almost abolished with only slightly different from the ∆CD1157 mutant under the same conditions tested. We did not observe a growth defect for any of the strains in anaerobiosis (Figure 50). We also tested the effect of these genes inactivation on the growth of C. difficile in TY soft agar tubes incubated in the presence of air for 24h. As control, we used the sigB mutant previously showed to be less tolerant to air (Kint et al. 2017). The ∆CD1474, ∆CD1524 and CD1157 single mutants and the ∆CD1474-∆CD1524 double mutant were able to grow as the wild-type strain (data not shown). By contrast, we observed a higher growth inhibition zone for the ∆CD1623 mutant compared to that of the wild-type strain (Figure 51A). The difference between the growth inhibition distance between the sigB and ∆CD1623 mutants and the wild-type strain was 6 and 4 mm, respectively (Figure 51B). The air sensitivity of the ∆CD1623-CD1157 double mutant will be addressed in future experiments.

complementation
Altogether these results indicate that the two FDPs, CD1157 and CD1623, and the two RevRbrs, CD1474 and CD1524, play an important role for the O 2 tolerance in C. difficile. We can also conclude that CD1474 and CD1524 are functionally redundant in their ability to reduce O 2 . Work is currently in progress in the laboratory to construct multi-mutants of both the FDPs and revRbrs genes and to complement the mutants inactivated with the respective genes.

P (sigB) -SNAP Cd

MTG TMR-Star Merge

II. Proteins produced under the control of σ B are present in the spores

In C. difficile, the alternative sigma factor σ B plays a pleiotropic and crucial role regulating numerous genes involved in biological processes such as motility, sporulation, cell wall metabolism and stress responses (Kint et al. 2017). However, while σ B positively controls stress response, the sporulation genes are negatively controlled by this sigma factor.

The majority of the genes of the σ F , σ E and σ G regulons (84%, 96% and 96%, respectively)

were up-regulated in the sigB mutant (Kint et al. 2017). However, only 38% of the σ K regulon was derepressed in the sigB mutant, a rate probably underestimated due to the timing of cell sampling used in the transcriptomic study (10 h) that did not allow detection of the expression of later sporulation genes controlled by σ B . In agreement, the inactivation of sigB resulted in a higher sporulation efficiency compared with the wild-type strain (Kint et al. 2017).

II.1. σ B expression and activity during sporulation

Most of the proteins located at the spore surface are encoded by genes transcribed in the MC by σ E and σ K . Interestingly, several proteomic studies revealed the presence of proteins at the spore surface that are not synthesized under the control of sporulation-specific sigma factors. In particular, the revRbrs, CD1474 and CD1524, the FDP CD1623 and the OR CD0176, has been identified in spore coat proteomes (Lawley et al. 2009b, Abhyankar et al. 2013, Diaz-Gonzalez et al. 2015, Kint et al. 2017). The corresponding genes are transcribed by σ B (Kint et al. 2017). Therefore, we decided to explore both the synthesis of σ B and its activity during sporulation. To monitor its expression during sporulation, the promoter region of the sigB operon located upstream of the CD0007 was fused to the SNAP Cd gene (Pereira et al. 2013, Cassona et al. 2016). The resulting plasmid was transferred into the 630∆erm strain.

After 24 h of growth in sporulation medium (SM), cells expressing the transcriptional P sigB -SNAP Cd fusion were doubly labeled with TMR-Star and the membrane dye MTG to allow identification of the different stages of sporulation. The expression of the P sigB -SNAP Cd fusion was detected in 63% of pre-divisional cells (Figure 52A). Moreover, fluorescence was detected in both MC and FS compartment after asymmetric division and during engulfment morphological steps in 54% and 28% of the cells, respectively (Figure 52A). Interestingly, at the end of sporulation after engulfment completion, the expression of the P sigB -SNAP Cd fusion was confined to the MC compartment, in 59% of the cells (Figure 52A). To check the activity of σ B along sporulation, we used the revRbr-encoding gene CD1474 as a σ B -target gene. Indeed, this gene is a good candidate to study σ B activity since a unique σ B -dependent promoter was mapped by TSS analysis upstream of the CD1474 gene, and its expression is completely abolished in the sigB mutant during vegetative growth (Soutourina et al. 2013, Kint et al. 2017) (Kint et al, in Annex). The promoter region of CD1474 was fused to the SNAP Cd gene and the resulting plasmid transferred into the 630∆erm strain and the sigB mutant. The expression of P CD1474 -SNAP Cd fusion was found in 81% of the pre-divisional cells but also in 86% of the cells after asymmetric division and in 75% of the cells during engulfment process in both MC and FS compartments. After engulfment completion, fluorescence signal was detected in 90% of the cells and again only in the MC compartment as observed for the P sigB -SNAP Cd fusion. In addition, no signal was detected in the sigB mutant (Figure 52B). Similar results were observed with the CD1524 and the CD1623 genes (Figure 53A andB). Interestingly, for the CD0176 gene, even if we observed the same compartmentalization of the σ B activity, the percentage of cells expressing P CD0176 -SNAP Cd fusion was much more low comparing with the other tested genes (Figure 53C). After engulfment completion, only 7% of the cells showed fluorescent signal. Importantly, for all the genes, no signal was detected in the sigB mutant strain. Altogether, these observations showed that both sigB and the σ B -controlled genes, CD1474, CD1524, CD1623 and CD0176 are expressed during sporulation and that this expression is surprisingly restricted to the MC compartment after engulfment completion. The absence of fluorescence in the sigB background indicates that even during sporulation, their expression is exclusively controlled by σ B .

II.2. Localization of σ B -controlled proteins at the spore surface

The detection of σ B activity in the MC during sporulation is in agreement with the identification of σ B -targets at the surface of the spore. Analysis of the ultrastructure of sigB mutant spores using TEM revealed that the sigB mutant produce spores similar to the wildtype strain, with intact spore coat and exosporium layers surrounding the cortex (Figure 54A).

However, in order to confirm the presence of σ B -target proteins at the spore surface, we compared the proteome of the spore surface layers of the wild-type and sigB mutant strains.

First, a SDS-PAGE analysis of the spore coat plus exosporium extracts obtained from purified spores did not reveal major differences in the protein profile of the sigB mutant compared with the wild-type strain (Figure 54B). To have a global view of the spore surface proteome of the sigB mutant, proteins extracted from the coat/exosporium of spores of the wild-type

CD1524-SNAP Cd

Results 120 strain and the mutant were analysed using mass spectrometry. This analysis revealed that the abundance of proteins previously found at the spore surface layers whose synthesis is under the control of σ B , decreased in the sigB mutant spores compared with the wild-type strain (Table 2). For instance, the abundance of the revRbrs CD1474 and CD1524 and the FDP CD1623 was around 8-, 3-and 4-fold reduced, respectively, in the sigB mutant. The abundance of the oxidoreductase CD0176 also decreased 7-fold in the coat extract of the sigB mutant. The nitroreductase CD1125, produced under the control of σ B , was also present at a reduced level in the coat extract of this mutant. In contrast, some proteins, such as the Rbr CD2845, were detected in a higher abundance in the coat extract of the sigB mutant compared to the wild-type strain. The CD2845 gene is controlled by the late FS sigma factor σ G (Table 2). Except for CD2845, we did not observe differences in the abundance of other spore coat proteins. We noted that contrary to other proteomic studies on C. difficile spores, we detected the FDP CD1157 in the coat extract of the wild-type strain. Moreover, no significant differences were observed in the abundance of this protein in the coat extract of the sigB mutant compared to the wild-type and strain (Table 2).

In order to confirm the localization of some of the proteins in the spore surface layers, we constructed C-terminal SNAP-tag translational fusions with some σ B -controlled genes.

These fusions were introduced into the wild-type and sigB mutant strains. Spores carrying the translational fusions were purified. The spore coat/exosporium was extracted in order to be analysed by Western blotting using an antibody raised against the SNAP protein. We have thus confirmed the presence of the revRbrs, CD1474 and CD1524, the FDP CD1623 and the OR CD0176 at the spore surface layers of the wild-type strain while these protein fusions were not detected in the coat/exosporium extract of the sigB mutant (Figure 55). The FDP CD1157 was detected in both wild-type and sigB mutant in agreement with the data of the proteome of the spore surface (Figure 55). However, a decreased amount of CD1157 was observed in the spore surface layers of the sigB mutant background in agreement with the expression of the CD1157 gene under the dual control of a σ A -and a σ B -dependent promoter.

Further experiments will be required in order to determine the role of these proteins in the spore and to decipher the molecular mechanisms involved to restrict the activity of σ B in the MC compartment at late stages of sporulation.

Materials and methods

Bacterial strains and growth conditions

The bacterial strains used in this study are listed in Table S1. E. coli strains were grown in LB broth (1% bacto-tryptone, 0.5% yeast extract, and 0.5% NaCl, pH 7.0) or in M9 minimal medium. When indicated, ampicillin (100 µg/ml), chloramphenicol (15 µg/ml) or kanamycin (30 µg/ml) was added to the culture medium. C. difficile strains were grown anaerobically (5% H 2 , 5% CO 2 and 90% N 2 ) at 37°C in brain heart infusion (BHI) medium (Difco), in TY (Bacto tryptone 30 g.l -1 , yeast extract 20 g.l -1 , pH 7.4) or in C. difficile minimal medium (CDMM). For solid media, agar was added to a final concentration of 17 g.l -1 .

Sporulation assays were performed in sporulation medium (SM) (Wilson et al. 1982). To produce spores, we used an SMC medium containing (per liter): 90 g of Bacto peptone (Bacto TM 211677), 5 g of Proteose peptone (82450, Sigma-Aldrich), 1 g of (NH 4 ) 2 SO 4 and 1.5 g of Tris. When necessary, cefoxitin (25 µg/ml), thiamphenicol (15 µg/ml) and erythromycin

(2.5 µg/ml) were added to the C. difficile cultures. All the primers used in this study are listed in Table S2.

Construction of C. difficile strains

The ∆CD2399, ∆CD1474, ∆CD1524 and ∆CD1623 knock-out mutants were obtained using the codA-mediated allele exchange method (Cartman et al. 2012, Peltier et al. 2015).

Briefly, for all these genes, 1.5 kb fragments located upstream and downstream of the gene to be deleted were PCR amplified from 630∆erm genomic DNA using primers CF14/CF15 and CF16/CF17, IMV917/NK70 and NK71/IMV918, IMV919/NK64 and NK65/IMV920, IMV914/NK58 and NK59/IMV915 (Table S2) for ∆CD2399, ∆CD1474, ∆CD1524 and ∆CD1623, respectively. Purified PCR fragments were introduced into the pMTLSC7315 plasmid using BamHI and XhoI restriction sites in case of ∆CD2399, and using the Gibson Assembly® Master Mix (Biolabs) for the other genes. The sequences of the resulting plasmids were verified by sequencing. The plasmids obtained were introduced in HB101/RP4

E. coli strain allowing the transfer by conjugation into the C. difficile 630∆erm strain.

Transconjugants were selected on BHI plates supplemented with Tm and Clostridium difficile selective supplement (SR0096, Oxoid). Isolation of faster growing single-crossover integrants was performed by serial restreaking on BHI plates supplemented with Cfx and Tm. Singlecrossover integrants were then restreaked on CDMM plates supplemented with fluorocytosine Results

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(50 µg/ml) allowing the isolation of double crossover events (Cartman et al. 2012). To confirm the plasmid loss, fluorocytosine resistant clones were restreaked on BHI plates supplemented with Cfx and Tm. The Tm sensitive clones were checked by PCR for the presence of the expected deletion (Figure S1). Steps in C. difficile were repeated in each different mutant in order to generate double-mutants, CDIP714 for the two revRbrs and CDIP1370 for the two FDPs (Table S1).

The ClosTron gene knockout system (Heap et al. 2007, Heap et al. 2009) was used to inactivate the CD1157 gene in 630∆erm, yielding strain 630∆erm CD1157::erm (CDIP588).

We designed with the Targetron design software (Sigma-Aldrich) primers to retarget the group II intron of pMTL0007 to insert into the CD1157 gene in sense orientation immediately after the 225 th nucleotide in the coding sequence (Table S2). The PCR product generated by overlap extension was cloned between the HindIII and BsrG1 sites of pMTL007 to obtain pDIA6374. E. coli HB101(RP4) containing pDIA6374 was mated with C. difficile 630∆erm.

C. difficile transconjugants were selected by sub-culturing on BHI agar containing Tm and Cfx and then plated on BHI agar containing Erm. Chromosomal DNA of transconjugants was isolated as previously described [START_REF] Antunes | CcpA-mediated repression of Clostridium difficile toxin gene expression[END_REF]. PCR was realized to verify the integration of the intron into the CD1157 gene and the splicing of the group I intron from the group II intron after integration (Figure S1).

To complement CD2399, CD1474 and CD1524, both the promoter region and the open reading frame of the corresponding gene were amplified by PCR using pairs of oligonucleotides, CF117/CF118, CF115/CF116, CF113/ CF114 for CD2399, CD1474 and CD1524, respectively. The PCR products were cloned into pMTL84121 to produce pDIA6534, pDIA6538 and pDIA6537 (Table S1). E. coli HB101 (RP4) strain containing each plasmid was mated with every single or double mutant (Table S1). For CD2399 complementation, the plasmid was transferred by conjugation into C. difficile ∆CD2399 (Table S1).

Bacterial adenylate cyclase two hybrid system (BACTH) and ßgalactosidase assays

Protein-protein interactions were analysed using the BACTH system (Euromedex). All the procedures for plasmid constructions and E. coli co-transformations were performed according to the manufacturer's instruction. Briefly, this system is based on the catalytic domain of the adenylate cyclase from Bordetella pertussis (CyA) that is composed of two

His-tag pull down assays

To construct the different plasmids, fragments were amplified from chromosomal DNA. For His-tagged protein construction, the fragments were cloned into petDuet using EcoRI and NotI restriction enzyme sites. For untagged proteins the fragments were cloned using Nde and XhoI (Table S2). Ligation products were transformed into NEB and confirmed by sequencing. The resulting plasmids were transformed into BL21 (DE3) cells for protein production. E. coli BL21 (DE3) strains (see Table S1) were grown to mid-log phase in 2YT ( 5g NaCl, 10 g yeast extract, and 15 g tryptone per liter), induced with IPTG, and grown overnight at 19°C. Cultures were pelleted, resuspended in lysis buffer (300 mM NaCl, 50 mM

Transcriptional SNAP Cd fusions

To construct SNAP transcriptional fusions, we used pFT47, a pMTL84121 plasmid derivative containing a synthetic SNAP Cd sequence (Pereira et al. 2013, Cassona et al. 2016).

To construct a sigB-SNAP Cd transcriptional fusion, pDIA6325 used for sigB complementation containing both the sequence located upstream of CD0007 (position -120 to +50 from the TSS) and the sigB gene was digested by XhoI and StuI to remove the sigB gene and keep the σ A -dependent promoter present upstream of CD0007. The SNAP Cd gene was PCR amplified using pFT47 DNA and primers pairs NK155/NK156. The DNA fragment was digested by DpnI to remove the pFT47 plasmid trace and inserted between the XhoI and StuI sites of pDIA6325 to obtain pDIA6589. The resulting plasmid was then transferred into the 630∆erm strain and the sigB mutant.

To construct the transcriptional SNAP Cd fusion with the complete promoter region of CD1157, a fragment (position -308 to +114 from the start codon) was amplified using genomic DNA and the primer pair IMV949/IMV950 to produce a 423 bp PCR product. The DNA fragment was inserted between the EcoRI and XhoI sites of pFT47 to obtain pDIA6517.

To construct the fusion bearing only the σ A -or the σ B -dependent promoter, promoters regions of CD1157 (position -308 to -64 and -104 to +114 from the start codon) were amplified with IMV949/IMV977 and IMV976/IMV950, respectively. The DNA fragments were then introduced between the EcoRI and XhoI sites of pFT47 yielding pDIA6670 and pDIA6669

for σ A -or the σ B -dependent promoter, respectively. To construct the CD1474-, the CD1524and the CD1623-transcriptional SNAP Cd fusions, the promoter regions (-206 to +19, -190 to +30 and -202 S1).

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Translational SNAP Cd fusions

To obtain C-terminal SNAP-tag translational fusions, we used pFT58 (Pereira et al. 2013, Cassona et al. 2016). pFT58 digested with EcoRI was amplified by inverse PCR using CF95/CF96. The promoter and coding regions of CD1474, CD1157 or CD1623 were amplified from genomic DNA using primer pairs CF99/CF100, CF107/CF108 and CF101/CF102. For CD0176-SNAP translational fusion, the promoter was amplified using CF103/CF104 and the CD0176 coding sequencing with CF105/CF106. The PCR products were inserted into pFT58 using Gibson Assembly yielding pDIA6497, pDIA6498, pDIA6499 and pDIA6503. The inserts of these plasmids were verified by sequencing. Plasmids were introduced in E. coli HB101 (RP4) and transferred to C. difficile 630∆erm and sigB mutant strains by conjugation (Table S1).

Fluorescence microscopy and image analysis

For SNAP labelling, the TMR-Star substrate (New England Biolabs) was added to the cells at a final concentration of 250 nM, and the mixture was incubated for 30 min in the dark.

After labelling, the cells were collected by centrifugation (4000xg for 3 min at room temperature), washed 3 times with 1 ml of PBS, and finally resuspended in 1 ml of PBS containing or lacking the membrane dye Mitotracker Green (MTG; 0.5 µg/ml) (Molecular Probes, Invitrogen). For phase-contrast and fluorescence microscopy, spores or cells were mounted on 1.7% agarose-coated glass slides to keep the cells immobilized and to obtain sharper images (Glaser et al., 1997). The images were taken with exposure times of 200 ms for MTG and 600 ms for SNAP. Cells were observed using a Nikon Eclipse TI-E microscope 60x Objective and captured with a CoolSNAP HQ2 Camera. The images were analysed using ImageJ.

Spore production and sporulation tests

SMC liquid medium inoculated with C. difficile was incubated overnight at 37°C. A sample (200 µl) of this culture was plated in SMC agar and incubated at 37°C for 7 days.

Spores scraped off with water were incubated for 7 days at 4°C. Cell fragments and spores were separated by centrifugation using a HistoDenz gradient (Sigma-Aldrich) (Dembek et al. 2013). Suspensions (>99% of spores) are free of vegetative cells, sporulating cells and cell debris as determined by phase-contrast microscopy. The spore suspension was quantified by measuring the OD 600nm .

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Overnight cultures grown at 37ºC in BHI were used to inoculate SM media at a dilution of 1:200. At specific time points, 1 ml of culture was withdrawn, serially diluted in BHI and plated before and after heat treatment (30 min, at 60ºC), in order to determine the total and heat-resistant colony forming units (CFU). Samples were plated onto BHI supplemented with 0.1% TA (Sigma-Aldrich) to promote efficient spore germination (Wilson et al. 1982).

Germination assay

Approximately 10 8 spores (equivalent of 0.4 OD 600nm units) were resuspended in 200 µl of water. The spore suspension was heat-treated at 65ºC during 10 min, and serial dilutions were plated on BHI containing 0.1% TA to determine the number of spores that are able to germinate. The CFU were monitored 24 h and 48 h after plating. At least three biological replicates were performed using at least two independent spore preparations. The germination of the spores was also monitored by OD 600nm (Donnelly et al. 2016) and by phase-contrast microscopy (Kong et al. 2011). Spores resuspended in BHI were exposed to 1% TA, and the OD 600nm was measured every 5 min for 1 h. For the phase-contrast microscopy, purified spores were resuspended to an OD 600nm of 0.4 in BHI containing 1% TA and incubated under anaerobic conditions for 4 h at 37°C. At each time point, samples were pelleted by centrifugation, washed with PBS and fixed with 2% paraformaldehyde for 15 min at room temperature. Fixed samples were washed with PBS once more and dried onto glass slides.

Phase-contrast microscopy was conducted using an Axioskop microscope (Zeiss) with a 100X oil immersion lens. The images were captured using an Axiocam 105 colour (Zeiss).

Spore resistance treatments

Lysozyme resistance of spores was measured by suspending 10 8 spores in 200 µl of PBS in the presence of 250 µg/ml of lysozyme. Ethanol resistance of spores was measured by resuspending the spores in 200 µl of 70% ethanol. Heat resistance of C. difficile spores was determined at 80°C. For each condition, spores were incubated for 1 h at 37°C with shaking (300 rpm), and the samples were plated onto BHI agar containing 0.1% TA and incubated anaerobically for 24 h at 37°C.

Transmission electron microscopy

To analyse the ultrastructure by TEM, purified spores were fixed with 2.5% of glutaraldehyde (Sigma Aldrich) in 0.1 M HEPES (4-(2-hydroxyethyl)-1piperazineethanesulfonic acid), pH 7.2. Post fixation was conducted with 1% osmium Results 127 tetroxide (Merck) and 1.5% ferro-cyanide (Sigma Aldrich) in 0.1 M HEPES. After dehydration by a graded series of ethanol, spores were infiltrated with epoxy resin. Seventynanometer sections were collected on formvar coated slot grids (EMS) and were contrasted with 4% uranyl acetate and Reynolds lead citrate. The stained sections were observed using a Tecnai spirit FEI operated at 120 kV. The images were acquired using an FEI Eagle digital camera.

Coat extraction and Western blot analysis

To analyse the profile of the coat and exosporium proteins, C. difficile spores were resuspended in extraction buffer (0.125 mM Tris-HCl, 5% β-mercaptoethanol, 2% SDS, 0.5 mM dithiothreitol (DTT) and 5% glycerol, pH 6.8) containing 0.025% bromophenol blue and boiled as previously described (Costa et al. 2006). For vegetative and sporulating cells, whole cell lysates were prepared as described previously (Fagan and Fairweather 2011). Briefly, sporulating cultures of C. difficile were harvested by centrifugation at 5000 rpm for 10 min at 4°C, and the pellets were frozen at -20°C. The bacteria were thawed and resuspended in PBS containing 0.12 µg/ml of DNase I to an OD 600nm of 40. An equal volume of 2X extraction buffer containing 0.025% bromophenol blue was added to the protein samples (Laemmli 1970), which were resolved by SDS-PAGE and visualized by Coomassie brilliant blue R-250 staining (Bio-Rad). Proteins were transferred to a PVDF membrane (0.45 µm) and subjected to immunoblot analyses as described previously (Serrano et al. 1999). Rabbit polyclonal

Mass spectrometry of the spore coat and exosporium layers

C. difficile spores were resuspended in extraction buffer and boiled as previously described (Costa et al. 2006). Coat and exosporium proteins were recovered by centrifugation and precipitated with 10% TCA. Protein pellets were resuspended in urea 8 M and NH 4 HCO 3 100 mM and sonicated on ice. Subsequently, 50 µg of total proteins were reduced with TCEP 10 mM (646547 -Sigma) for 30 min and then alkylated with iodoacetamide 20 mM (I114 -Results

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Sigma) for 1 h. Proteins were digested with rLys-C 1 µg (V1671, Promega) for 4 h at 37°C

and then digested with trypsin 1 µg (V5111, Promega) overnight at 37°C. The digestion was terminated with formic acid 4%, and the peptides were desalted using a C18 Sep-Pak Cartridge (WAT054955, Waters). The peptides were eluted successively with 50% Acetonitrile (ACN)/0.1% Formic acid (FA) and 80% ACN/0.1% FA before being dried in a vacuum centrifuge. The peptides were resuspended in 2% ACN /0.1% FA.

For LC-MS/MS analysis, a nanochromatographic system (Proxeon EASY-nLC 1000, Thermo Fisher Scientific) was coupled online to a Q Exactive TM HF Mass Spectrometer (Thermo Fisher Scientific). For each sample, 1 µg of peptides was injected onto a 50-cm homemade C18 column (1.9 µm particles, 100 Å pore size, ReproSil-Pur Basic C18, Dr.

Maisch GmbH) and separated with a multi-step gradient from 2 to 45% ACN in 105 min at a flow rate of 250 nl/min over 132 min. The column temperature was set to 60°C. The data were acquired as previously described (Lago et al. 2017).

Raw data were analyzed using MaxQuant software version 1.5.1.2 (Cox and Mann 2008) and the Andromeda search engine (Cox et al. 2011). The MS/MS spectra were searched against an internal C. difficile database containing 3 957 proteins, and the contaminant file included in MaxQuant. The database search parameters were fixed as described (Lago et al. 2017). Output proteinGroup file was integrated into Perseus [START_REF] Tyanova | The Perseus computational platform for comprehensive analysis of (prote)omics data[END_REF] to perform data filtering and statistical tests. First, contaminants, reverse identifications and proteins only identified by site were excluded. A protein filtering was set for the validation process such as a protein was integrated in the final list only if this one was identified in at least two replicates of a condition. Second, LFQ intensities were log2 transformed and missing values were imputed and replaced by random Log2 LFQ intensities that are drawn from a normal distribution at the low detection level. Third, a two-sided T-tests of the Log transformed LFQ intensities with a permutation-based FDR calculation at 5% and S0=0.5 [START_REF] Tusher | Significance analysis of microarrays applied to the ionizing radiation response[END_REF] were applied to determine statistically significant proteins.

Protein expression and purification

The coding region of CD1474 and CD1524 were PCR amplified using genomic DNA and primer pairs IMV970/IMV971 and IMV968/IMV969 to produce a 540 bp PCR product.

The resulting PCR products were cloned into pET20 (Novagen) yielding pDIA6635 and pDIA6671. The amino acid sequence of CD1157 was used to synthesize its encoding gene at GenScript Inc. USA, codon optimized for expression in E. coli. The sequence was cloned into pET24a (Novagen). After verification by sequencing, the plasmids were introduced in E. coli Results 129 BL21 (DE3). These strains were used for overexpression of the respective proteins.

Overexpression of CD1524, CD1474 and CD1157 was performed by growing the strains aerobically at 37 °C at 150 rpm in M9 minimal medium, with 20 mM glucose, 0.1 mM FeSO 4 , and 100 µg/ml ampicillin (for CD1474 and CD1524), or 50 µg/ml kanamycin (for CD1157) until an optical density at 600 nm of approximately 0.4 was reached. At that time, 0.1 mM of FeSO 4 was added and protein expression was induced either by 1 mM of IPTG for CD1524 and CD1474 or 100 µM for CD1157. After 6 h of growth at 30°C, cells were harvested by centrifugation, resuspended in a buffer containing 20 mM (Tris)-HCl (pH 7.5) and stored at -20 °C. Cells were disrupted by at least 3 cycles in a French Press apparatus at 16.000 psi (Thermo) in the presence of DNase (Applichem). The crude extract was cleared by low-speed centrifugation at 25.000xg for 25 min and then at 138.000xg for 1 h 30 min at 4 °C to remove cell debris and membrane fraction, respectively. The soluble extract was dialyzed overnight at 4 °C against buffer A (20 mM Tris-HCl, pH 7.5, 18% Glycerol) and was subsequently loaded onto a Q-Sepharose Fast Flow column (65 mL, GE Healthcare) previously equilibrated with buffer A. Proteins were eluted with a linear gradient from buffer A to buffer B (buffer A containing 500 mM NaCl). The eluted fractions were followed throughout the purification process by SDS-PAGE and UV-visible spectroscopy. Fractions containing the desired protein were pooled and concentrated. For CD1474 and CD1524, the concentrated fraction was then loaded onto a size exclusion Superdex S75 column (330 ml, GE Healthcare), equilibrated with buffer A containing 150 mM NaCl. The fractions containing the protein were pooled and concentrated. For the CD1474, the last fraction was loaded onto a Fractogel column (20 ml, Merck Millipore), previously equilibrated with buffer

A and eluted with a linear gradient from buffer A to buffer B. After purification, CD1157 was incubated overnight at 4°C in the presence of 1 mM of flavin mononucleotide (FMN) and subsequently applied onto a PD-10 desalting column to eliminate unbounded FMN. Fractions containing purified proteins were verified by SDS-PAGE (Figure 44).

Protein, metal and flavin quantification

Purified protein samples were quantified using the Bicinchoninic Acid Kit (Thermo)

and bovine serum albumin as standard. Iron content was determined by the phenanthroline colorimetric method. Protein samples were incubated for 15 min with 1 M HCl and for further 30 min with 10% trichloroacetic acid at room temperature, centrifuged at 5,000 g for 20 min and neutralized with the addition of 15% of ammonium acetate. Samples were then incubated Results 130 with 10% hydroxylamine and 0.3% 1-10-phenanthroline. The absorbance spectrum was measured, and iron content quantified using ɛ 510nm = 11.2 mM -1 .cm -1 . The flavin type was analysed as previously described (Folgosa et al. 2018b).

Oligomeric State Determination

The oligomeric state of the proteins was determined by size exclusion chromatography. Proteins were loaded onto a 25 ml Superdex S200 10/300 GL column (GE Healthcare), previously equilibrated with buffer A containing 150 mM NaCl. We used as standards a mixture containing aldolase (158 kDa), ferritin (440 kDa), conalbumin (75 kDa), carbonic anhydrase (29 kDa) and cytochrome c (12.4kDa), and dextran blue (2000 kDa) as void volume marker.

Redox titrations

The diiron center of CD1157, CD1524 and CD1474, and the corresponding reduction

Oxygen tolerance assays

O 2 tolerance assays were performed as indicated below. Five microliters of different serial dilutions (from 10 0 to 10 -5 ) of C. difficile strains grown for 7 h in the TY medium were spotted on TY plates supplemented with 0.05% of taurocholate. Plates were then incubated in the presence of different O 2 tensions while a control plate was incubated in anaerobiosis. The last dilution allowing growth was recorded after incubation at 37°C for 64 h.

Spectrophotometric measurements of the H 2 O 2 reductase activity

Because the physiological electron donor to the C. difficile CD1524, CD1474 and CD1157 is unknown, we used an artificial electron-donating system: a mixture of a NADH:rubredoxin oxidoreductase (NROR) and a rubredoxin from E. coli. The E. coli proteins were purified as previously described [START_REF] Gomes | Spectroscopic studies and characterization of a novel electron-transfer chain from Escherichia coli involving a flavorubredoxin and its flavoprotein reductase partner[END_REF][START_REF] Vicente | Kinetics of electron transfer from NADH to the Escherichia coli nitric oxide reductase flavorubredoxin[END_REF]). The enzymatic activity for H 2 O 2 was determined by UV-Visible spectroscopy, inside an anaerobic chamber (Coy Lab Products). The assays were performed in buffer C. The reaction was monitored at 340 nm, observing the NADH consumption (ε 340 = 6220 mM -1 cm -1 ). A mixture of buffer, 200 µM of NADH, NROR, rubredoxin and each protein of interest was prepared inside a cuvette, after which the reaction was initiated by the addition of H 2 O 2 . Concentrations of CD1524, CD1474 or CD1157 were 0.03 µM, 0.1 µM and 1 µM respectively.

Measurement of O 2 reductase activity

The O 2 reductase activity of the proteins was measured amperometrically with Clarktype electrodes selective for O 2 (Oxygraph-2K, Oroboros Instruments, Innsbruck, Austria).

The assays were performed in buffer C. The O 2 reductase activity was evaluated at 25 °C in air equilibrated buffer (~250 µM of O 2 ), in the presence of 5 mM NADH, NROR (810 nM)

and rubredoxin (5 µM) from E. coli. The reaction was initiated by addition of CD1524 or CD1474 (0.1 µM) or CD1157 (1 µM). Assays were performed in the presence of catalase (640 nM) and superoxide dismutase (240 nM).

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Supplementary tables and figures

Table S1. Bacterial strains and plasmids used in this study. The erm gene encodes the erythromycine resistance cassette. The cat gene encodes the chloramphenicol resistance cassette leading to resistance to chloramphenicol, Cm R and to thiamphenicol, Tm R . Amp R and Kan R means that the plasmid harbors a resistance cassette responsible for ampicillin and kanamycin resistance, respectively. For the CD1157 Clostron mutant, the verification of the L1.LtrB intron integration into the CD1157 gene was performed by using primers IMV844/IMV845 flanking the insertion site in CD1157. For the ACE mutants, the verification of the deletion of CD1623, CD1474 and CD1524 genes in the different backgrounds was performing using CF24/CF31, CF19/CF30 and CF23/CF39 respectively.

Strains

Genotypes

-(CD1157) EcoRI IMV950 CCGCTCGAGAGTTGTAAGTTGTCCCAA 3' Pσ B -(CD1157) XhoI IMV968 GGAATTCCATATGAAGAAATTTGTTTGTACAG 5' CD1474 pET20 NdeI IMV969 CCGCTCGAGTTATTATTTTCCAAAATATCTATTTAATAA TC 3' CD1474 pET20 XhoI (stop) IMV970 GGAATTCCATATGAAAAAATTTGTTTGTACAG 5' CD1524 pET20 NdeI IMV971 CCGCTCGAGTTATTATTTTCCAAAATGTCTGTTTAA 3' CD1524 pET20 XhoI (stop) IMV976 AGGAATTCATAAAACTATTGTTTTACATGA 5' Pσ B -(CD1157) EcoRI IMV977 CCGCTCGAGAAAAGATGTCATGATAAGTCA 3' Pσ A -(CD1157) XhoI NK58 GTTATTTAGTCTCTTTATACCCTGCTTG ACE CD1623 3' PCR1 NK59 CAAGCAGGGTATAAAGAGACTAAATAACGATTTTGAGG ATATTATGAAGGCTG ACE CD1623 5' PCR2 rev comp NK58 NK64 GGTGCAGCATCTCCTTCATGTATA ACE CD1524 3' PCR1 NK65 TATACATGAAGGAGATGCTGCACCCTTAGGATTATTAA ACAGACATTTTGG ACE CD1524 5' PCR2 rev comp NK64 NK70 ATTTCTTCATAATTACTTTCCTCCTAAA ACE CD1474 3' PCR1 NK71 TTTAGGAGGAAAGTAATTATGAAGAAAGAAGCTAGACA TGGTAAAGCATTC ACE CD1474 5' PCR2 rev comp NK70 NK155 GATCAGGCCTATGGATAAAGATTGTGAAATGA pFT47 5' StuI NK156 GATCCTCGAGTTATTCCTTACCCAAGTCCT pFT47 3' XhoI 1 1 1 1 1 1 2 2 2 2 2 2 3 3 3 3 3 3 4 4 4 4 4 4 CD1157 Wild-type ∆CD1623 ∆CD1623-CD1157 ∆CD1474- ∆1524 ∆CD1474-∆1524- CD1157 1: CD1157 : IMV844/

Discussion

I. The morphogenesis of the C. difficile spore coat

Spores play a key role in the pathogenesis of C. difficile. This metabolically inactive structure allows transmission, and its accumulation inside the host ensures persistence and recurrence. The anatomy of the spore comprises a series of specialized concentric shells, each of which provides specific functions [START_REF] Setlow | Spores of Bacillus subtilis: their resistance to and killing by radiation, heat and chemicals[END_REF][START_REF] Setlow | Spore Resistance Properties[END_REF]. A critical protective structure is the coat, a proteinaceous layer that encases bacilli and clostridia spores and provides a barrier against degradative enzymes and chemical compounds [START_REF] Nicholson | Resistance of Bacillus endospores to extreme terrestrial and extraterrestrial environments[END_REF][START_REF] Riesenman | Role of the spore coat layers in Bacillus subtilis spore resistance to hydrogen peroxide, artificial UV-C, UV-B, and solar UV radiation[END_REF][START_REF] Klobutcher | The Bacillus subtilis spore coat provides "eat resistance" during phagocytic predation by the protozoan Tetrahymena thermophila[END_REF][START_REF] Setlow | Spore Resistance Properties[END_REF]). Unlike B. subtilis, C. difficile possesses an exosporium, which is the outermost spore layer. The exosporium provides the first site of interaction with the host. Several proteins have been identified in the spore coat and exosporium layers of C. difficile and the function of some of them was already studied (Permpoonpattana et al. 2011b, Abhyankar et al. 2013, Permpoonpattana et al. 2013, Diaz-Gonzalez et al. 2015). However, additional studies are still needed to define the proteins and the mechanisms that guide deposition of these components in the outermost layers of C.

difficile spores and the interactions between them. Here, we have elucidated a new protein, CotL that controls the assembly of the spore coat layers in C. difficile.

I.1. Role of CotL on C. difficile spore coat structure

SpoIVA (Putnam et al. 2013) and SpoVM (Ribis et al. 2017), are the only conserved morphogenetic proteins between B. subtilis and C. difficile. In this study, we showed that the C. difficile coat-associated protein, CotL, is a new morphogenetic protein essential for the efficient assembly of the coat layers. Indeed, the assembly of early-and late-coat proteins occurs in a CotL-dependent manner and CotL is required for the appearance of the spore coat.

Consistent with this, the amount of most of the spore coat and exosporium proteins is strongly decreased in the spore surface layers of the cotL mutant when compared to the wild-type strain. However, cotL inactivation impaired neither the transcription of sporulation-related genes, nor the protein production in sporulating cells. Thus, we have concluded that CotL plays a morphogenetic rather than a regulatory role in coat formation.

I.2. The dual control of cotL gene expression

Our data shows that the CotL protein resides within the outermost layers of the spore, in agreement with the expression of cotL gene in the MC compartment under the dual control Discussion 142 of σ E and σ K . The cotL gene is transcribed by σ E early during sporulation suggesting a potential role of CotL in early FS development, as already observed for SpoIVA and SipL (Putnam et al. 2013). Then, the transcription of the cotL gene by σ K probably allows the maintenance or an increased expression of this gene that might be needed during the late sporulation stages. The timing of cotL expression has consequences on the assembly of CotL itself around the FS. Using SNAP translational fusions, we observed that when CotL is produced only under the control of σ K leading to a delayed and likely a reduced amount of its production, CotL seems to be immediately assembled at the FS surface. By contrast, when produced under the dual control of σ E and σ K , CotL first accumulates in the MC and then assembles around the FS. Moreover, the CotL protein produced under the control of its native promoter is not assembled around the FS in the sigK mutant. It has been shown that synthesis of coat proteins is usually controlled either by σ E or by σ K , but it is only when σ K becomes active that the coat layers start gaining their characteristic appearance as viewed by electron microscopy (Henriques and Moran 2007). Consistent with this, the sigK mutant itself is defective in coat polymerization and localization (Pereira et al. 2013, Pishdadian et al. 2015).

However, it has been shown that the CotB-SNAP Cd protein (a σ E -target) is located at the FS surface in the sigK mutant (Pereira et al. 2013), suggesting that C. difficile coat assembly can start at early sporulation stage as in B. subtilis (Henriques and Moran 2007). Our observations raised the hypothesis that CotL assembly might be dependent on the presence of a still uncharacterized σ K -regulated gene product even if we cannot exclude that the reduced amount of CotL when expressed solely under the control of σ K might be directly assembled without accumulation in the MC. The analysis of the CotL localization in different mutants inactivated for coat proteins is an interesting point to address in future work.

A continuous expression of cotL gene during sporulation suggests that the timing of CotL production may have an impact in the assembly of the other coat components around the FS. B. subtilis CotE, the outer coat morphogenetic protein located at the interface of the inner and outer spore coat layers, is an example of how the timing of protein synthesis is essential to determine a correct morphogenesis of the spore. As for C. difficile cotL, the cotE gene of B.

subtilis is similarly expressed first from a σ E -dependent promoter (P1) and later from a σ Kdependent promoter (P2) (Zheng and Losick 1990). The P1 promoter is shut off by SpoIIID at the early stages of the morphogenesis while the P2 promoter is shut off by GerE during the latest stages of morphogenesis. The cotE mutant retains the inner coat but lacks the outer coat layer (Zheng et al. 1988). Additionally, the early expression of cotE is essential for outer coat assembly since a later expression of this gene under the control of the σ K -dependent (gerE) or GerE-dependent (cotG) promoter, respectively does not allow assembly of a visible outer coat structure. In our case, we observed that C. difficile CotL is required for the correct encasement of the basement layer protein SpoIVA around the FS suggesting that the early synthesis of CotL is likely essential for the assembly of the C. difficile spore layers. How does CotL localizes in spoIVA mutant sporulating cells is an information that would give clues about its role in spore coat assembly.

I.3. CotL properties and its localization in the spore

The previous proteomic studies (Lawley et al. 2009b, Abhyankar et al. 2013, Diaz-Gonzalez et al. 2015) in addition to our proteomic analysis failed to detect the CotL protein in the spore or in the spore surface layers. However, we successfully detected CotL in the coat/exosporium extract of C. difficile spores by Western blot analysis using an antibody raised against CotL. The feature of CotL as a lysine-rich protein might hinder its detection by bottom-up mass spectrometry-based proteomics. Indeed, proteins are usually digested into peptides with trypsin, an enzyme that cleaves after lysine and arginine residues and the generated peptides of CotL digestion might be therefore too small or not suitable with their further identification by mass spectrometry.

The exact location of CotL within these spore layers still remains to be determined more precisely. In C. difficile, coat assembly around the FS is driven by SpoIVA and SipL (Putnam et al. 2013). In the corresponding mutants, the sporulating cells exhibit defects in coat localization, spore coat swirls are detected in the MC cytoplasm, and the spores cannot be purified (Putnam et al. 2013). CdeC and CdeM were identified as morphogenetic factors of the exosporium (Barra-Carrasco et al. 2013, Calderon-Romero et al. 2018). Spores of the cdeC mutant, but not that of cdeM mutant, exhibit a thinner coat (Calderon-Romero et al.

2018).

Our results indicate that the defect in spore formation of cotL mutant is less severe than that of spoIVA or sipL mutant but is more severe when compared to that of the cdeC mutant. We can purify cotL mutant spores, even if most of the coat and the exosporium layers the P σ E /σ K (cotL) -cotL-SNAP Cd was detected in the MC compartment (90% and 87%, respectively), around the spore in both the sporulating cells (8% and 7%) and in the free spores (2% and 6%) (Figure 56A). In contrast, the P σ K (cotL) -cotL-SNAP Cd fluorescence signal was detected either as a cap in the MCP pole (60% and 33%) or around the spore in both the sporulating cells (27% and 39%) and the free spores (13% and 26%) (Figure 56B). In agreement with our hypothesis, this data showed that CotL localization is not affected in the absence of CdeM (Figure 56). As mentioned above, how the inactivation of spoVM, spoIVA, sipL and/or cdeC genes influences the pattern of CotL localization is an interesting point to address in future work. Comparing with the B. subtilis model, our results suggest that C. difficile CotL acts earlier in coat morphogenesis than SafA and CotE. The safA mutant spores lack an inner coat but still retain the outer coat (Takamatsu et al. 1999). However, the cotE mutant spores lack an outer coat and retain the inner coat (Zheng et al. 1988), while the basement layer does not properly adhere to the underlying cortex (Henriques and Moran 2007). In C. difficile, few coat materials are detected around the purified spores of the cotL mutant and no lamellar or electrodense layers are observed, suggesting a stronger impact of the inactivation of cotL on spore morphogenesis than inactivation of the safA or the cotE genes in B. subtilis. Consistent with the observation that SpoIVA is not correctly assembled around FS in the cotL mutant, we proposed that CotL is an important player in starting coat assembly.

I.4. Interactions between coat proteins in C. difficile

The spore coat assembly requires a complex protein interaction network (Kim et al. 2006, McKenney andEichenberger 2012). Morphogenetic proteins make an initial structure from which the coat grown around the spore by the polymerization of individual proteins and addition of other components. In C. difficile, very little is known about interaction between spore coat proteins. So far, interactions between SpoIVA and SpoVM or SipL proteins were the only ones demonstrated (Putnam et al. 2013, Ribis et al. 2017). However, the binding of other coat proteins to SpoIVA and/or SipL remains unknown. The requirement of CotL for the encasement of other proteins at the spore outermost layers lead us to test possible interactions between CotL and coat and/or exosporium proteins such as SipL, CotA, CotB, CotE and CdeC. From all proteins tested, our experiments using the bacterial two hybrid system (BACTH) only suggested that CotL might interact with SipL. However, we were unable to confirm this result using pull down assays. In collaboration with Aimee Shen's group, we also tried to pull down CotL using a SipL-3xFlag fusion in C. difficile but we did not have more success. Concerning CotB and CotE, the interactions with CotL were weak and detected in one out of eight combinations only when CotL was fused to the T25 part of the adenylate cyclase through its C-terminal part. Whether in the other constructions, CotL contact regions might be blocked by the adenylate cyclase domains is a question that remains to be adressed. However, none positive confirmation of possible interactions came from the pull down assays. In addition, no interaction was revealed by the BACTH analysis between CotL and CotA or CdeC suggesting an absence of interaction. However, we cannot exclude that CotA or CdeC fusions were poorly synthethized, or that their stability and/or solubility were low in E. coli. Fail attempts to overexpress coat proteins in E. coli have been already reported [START_REF] Krajcikova | Exploring the interaction network of the Bacillus subtilis outer coat and crust proteins[END_REF]. A CotG self-interaction and a direct contact between CotG and CotB from B. subtilis was detected using a yeast two hybrid system [START_REF] Zilhao | Interactions among CotB, CotG, and CotH during assembly of the Bacillus subtilis spore coat[END_REF] while no interaction was revealed for CotG by BACTH analysis. It seems that overexpression of cotG in E. coli failed, explaining the lack of any observable CotG interaction using this system. Difficulties in over-expressing exosporium proteins under soluble conditions have been also reported. For instance, the C. difficile exosporium protein, CdeC, requires E. coli strains providing an oxidative environment for optimal soluble expression conditions [START_REF] Brito-Silva | Identification of Escherichia coli strains for the heterologous overexpression of soluble Clostridium difficile exosporium proteins[END_REF]). In addition, false positive and negative results can be generated by the BACTH system, which requires that verification must be done using other methods. As we observed for SipL, even if interactions were detected in E. coli, it might not mean that they indeed exist at the spore surface. Used as control, SipL and SpoIVA were shown to interact using pull down assays (Putnam et al. 2013) and we were able to reproduce this result.

Unfortunatelly, we were unable to made the SpoIVA constructs to show an interaction between SpoIVA and SipL by BACTH. To address and facilitate the screening of coat protein-protein interactions additional tools are required. Since bacterial-and yeast-two hydrid assays can give different results, we need in the future to optimize the overproduction and solubilization of coat/exosporium proteins in each organism. In C. difficile, the identification of partners of CotL might be possible by using a CotL-3xFlag fusion.

I.5. Spore encasement dependent on CotL

We showed that encasement of early-and late-proteins around the FS occurs in a CotL-dependent manner. However, no direct interaction between CotL and all the analysed proteins has been demonstrated so far. In B. subtilis, a large group of outer coat proteins requires CotE for localization. However, only a very few such as CotC, CotU and CotH interacts directly with CotE [START_REF] Isticato | CotE binds to CotC and CotU and mediates their interaction during spore coat formation in Bacillus subtilis[END_REF][START_REF] Isticato | The Direct Interaction between Two Morphogenetic Proteins Is Essential for Spore Coat Formation in Bacillus subtilis[END_REF]. In C. difficile, CotB, CotE and CdeC might be part of the CotL network, which does not necessarily require direct interaction. In fact, we cannot rule out the possibility that CotL might need the assistance of another unidentified factor essential for the physical contact between these proteins.

Additionaly, an interesting point in CotL protein is its amino acid composition. The positively charged amino acid, lysine, correspond to 24% of the CotL residues. Among the spore coat proteins of B. subtilis, the outer coat component, CotO, contains 16% of lysine residues.

Interestingly, CotO is a morphogenetic protein, which assembly is CotE-dependent. CotO directs the deposition of a subset of coat proteins and is needed to form a morphologically normal outer coat [START_REF] Mcpherson | Characterization of the Bacillus subtilis spore morphogenetic coat protein CotO[END_REF]. Whether the high percentage of lysines is important for the function of CotO in B. subtilis has not been addressed. However, the high level of this charged amino acid might be relevant for the assembly of the coat layers. At physiological pH, we deduce that C. difficile CotL is positively charged (pI 9.76) while most of the spore coat proteins are negatively charged. This unique propertie of CotL might be relevant to mediate electrostatic interactions with other structural components and could contribute to the formation of a tough layer and the mechanical resistance of the spore coat.

Since we found by BACTH positive interactions between CotL and SipL, and to a lesser extent with CotB or CotE, if these interactions are mainly electrostatic salt concentration and pH are probably important factors to take into account to optimize the pull down assays.

I.6. Model for the assembly of the outermost layers of the C. difficile spore

Altogether, our results allow to elaborate a preliminary model for the assembly and organization of the outermost layers, coat and exosporium in C. difficile spores (Figure 57A).

In a basement layer, SpoIVA and SipL interact and might form a complex that drive coat assembly around the FS (Putnam et al. 2013). Similar to B. subtilis, SpoVM probably recognizes the positive curvature of the FS and embed itself within the MC-derived membrane. Since SpoVM is largely dispensable for SpoIVA encasement in the C. difficile FS, SpoIVA may independently interact with the FS membrane (Ribis et al. 2017). Alternatively, SipL or another factor could help SpoIVA to surround the FS in the absence of SpoVM (Figure 57A). TEM allowed to distinguish an inner lammelar and an outer electrodense coat in C. difficile spores (Permpoonpattana et al. 2011b, Barra-Carrasco et al. 2013). However, it remains unclear whether each distint layers are associated to specific morphogenetic proteins The morphogenetic proteins of the exosporium, CdeC and CdeM, direct the assembly of the most external layer (Calderon-Romero et al. 2018). CdeC might be at the interface of the spore coat and exosporium layers. However, CdeM seems to be located uniquely in the exosporium layer (Figure 57A). Coupling our findings with previous localization studies (Diaz-Gonzalez et al. 2015), we can propose putative locations for the other spore surface proteins identified (Figure 57B). CotA and CotB are located in the spore coat layers (Diaz-Gonzalez et al. 2015) while the CotE, CotD, BclA and Cde proteins are located in the exosporium (Pizarro-Guajardo et al. 2014). The CotL-dependent proteins, defined as those that were mis-assembled in the cotL mutant FS, include SpoIVA, CotB, CotD, CotE and CdeC (Figure 57B) and cannot be precisely located. In fact, the mis-assembly of early coat proteins in the cotL mutant, most likely hinders the correct localization of late coat proteins such as CdeC, CotD and CotE and consequently the assembly of the exosporium layer.

Finally CdeC is required for the assembly of CdeA, CotB and the high molecular complex BclA1 and BclA3, while CdeM-dependent proteins include CotA and CdeB, the low molecular mass complex formed by BclA1, BclA2, BclA3 and CdeB (Figure 57B). ??

II. Signaling pathway and σ B activation in C. difficile

Bacterial cells respond to stress or starvation by expressing a set of genes whose products are necessary to cope with the adverse environmental conditions and ensure survival of the cell [START_REF] Kjelleberg | How do non-differentiating bacteria adapt to starvation?[END_REF]). In C. difficile, the alternative sigma factor σ B controls genes involved in stress response (Kint et al. 2017). σ B equips the cells with a protective and preventive machinery conferring stress resistance (Hecker et al. 2007, Kint et al. 2017).

During my thesis, I participated with Nicolas Kint in the study of the signaling pathway leading to the activation of σ B in C. difficile. My main contribution in this work was in the analysis of both sigB expression and σ B activity during vegetative growth (Kint et al, in Annex).

II.1. Role of the environmental signals in the activation of σ B in C. difficile

In this study, Nicolas Kint showed the conservation of the partner switching mechanism, involving RsbV and RsbW, in the activation of σ B in C. difficile. He also demonstrated that the phosphorylation state of RsbV, and later on the activation of σ B , is dependent on the activity of the newly identified PP2C phosphatase CD2685/RsbZ (Figure 58). However, stress signals triggering phosphatase activation in C. difficile remain unknown.

In addition, no typical environmental stresses tested induce σ B activity. Indeed, exposure to bile salts, acidic pH, ROS or NO-donor compounds that correspond to stresses encountered within the gut did not increase the expression of σ B -target genes. When we tested a combination of NO-donor compounds with H 2 O 2 or a superoxide donor, mimicking the conditions encountered by C. difficile during inflammation, we did not observed induction of

II.2. Role of the energy signals in the activation of σ B in C. difficile

As in other Firmicutes, the activity of σ B in C. difficile increases at the onset of the stationary phase (Kint et al. 2017) and several σ B -targets are more expressed at 10h compared to 4h (Saujet et al. 2011). At the entry in stationary phase, energy deficiency occurs in consequence of the decreased intracellular concentration of ATP. In B. subtilis or L.

monocytogenes, the addition of ionophore carbonyl cyanide m-chlorophenylhydrazone (CCCP) leading to a drop of the ATP pool as well as the growth in starvation conditions of glucose, phosphate or in limitation in O 2 availability that likely result in ATP drop also allows the activation of σ B (Hecker et al. 2007). Nicolas Kint showed that the addition of CCCP induces the activity of σ B in C. difficile in a RbsZ-dependent manner (Figure 58). Since exposure to CCCP and entry into stationary phase are able to trigger σ B target expression, it is quite possible that σ B activation is mediated by an energy stress (Figure 58). Additional experiments to decipher the mechanism leading to the σ B activation in response to CCCP are currently being performed in the laboratory.

From a physiologic point of view, the activation of σ B through an energetic pathway raises some questions. Studies have shown that in dysbiotic mice there is an increase in the concentration of compounds required for growth and multiplication of vegetative cells of C.

difficile (Theriot et al. 2014, Theriot andYoung 2014). These compounds include amino acids such as glycine, cysteine or proline, as well as mannitol and sorbitol (Theriot et al. 2014).

These

II.3. Expression of sigB in C. difficile

In most Firmicutes, a basal expression of the sigB operon is mediated by a promoter recognized by σ A . σ B is produced at a basal level, sequestrated by RsbW and is eventually turned active following stress. However, the presence of an internal promoter upstream of the rsbV gene recognized by σ B promotes a rapid increase in the production of RsbV, RsbW and σ B when the bacteria are exposed to a stress (Hecker et al. 2007). Contrary to the other Firmicutes, σ B of C. difficile is not auto-regulated but only constitutively expressed throughout the growth via a σ A -dependent promoter. In agreement, we demonstrated by qRT-PCR and monitoring the expression of a P sigB -SNAP Cd fusion, that sigB gene is expressed similarly in the wild-type and the sigB mutant strains during growth. In B. subtilis and L.

monocytogenes, the last gene of the sigB operon, rsbX, codes for a PP2C phosphatase that allows the "reset" of the stressosome, decreasing the activity of σ B . This mechanism prevents excessive activation of σ B , which can be toxic to the bacteria [START_REF] Eymann | In vivo phosphorylation patterns of key stressosome proteins define a second feedback loop that limits activation of Bacillus subtilis sigmaB[END_REF], Xia et al. 2015). Similarly, in B. cereus, a mechanism mediated by RsbK and RsbM modulates the activity of σ B , preventing its permanent activation (Chen et al. 2012). None of the genes encoding these proteins are present in the C. difficile genome, suggesting that none of these systems exists for σ B in this bacterium. These results are consistent with the fact that the overexpression of rsbZ causes a growth defect and that overexpression of sigB is toxic in C.

difficile. Therefore, the absence of an auto-regulation of σ B in C. difficile might be a mechanism used by this bacterium to prevent over-activation of σ B , potentially toxic to the cell.

In addition, the fact that the overexpression of the rsbZ gene induces a stronger activation of σ B , suggests that some phosphatase might be active even in the absence of a stress. Interestingly, during exponential growth, a fraction of the total RsbV is dephosphorylated in C. difficile (Kint et al, in Annex) as observed in S. aureus and B. subtilis (Eymann andHecker 2001, Pane-Farre et al. 2009). It seems possible that RsbZ has a basal activity allowing the dephosphorylation of RsbV~P and therefore the activation of σ B in the absence of a stress exposure. This hypothesis is in agreement with our data obtained from SNAP fusions with genes strictly dependent on σ B for their expression, showing that σ B is active during the exponential growth phase at least in a fraction of the cells as already observed in S. aureus and L. monocytogenes (Pane-Farre et al. 2009, Utratna et al. 2012, Guldimann et al. 2017). The activation of σ B during the exponential growth phase without previous exposure to stress probably promotes a rapid adaptation of the bacterium when it copes a hostile environment. Under these conditions, early activation of σ B could be essential during the infectious cycle to resist the various stresses encountered in the host. However, we also observed that the revRbr CD1474 gene expression level increases during growth, confirming an increased σ B activity at the stationary phase, as observed in other Firmicutes, and in previous transcriptomic data of C. difficile comparing the expression profiles between exponential and stationary growth phase (Saujet et al. 2011).

II.4. Heterogeneity of gene expression of sigB and σ B -targets

Interestingly, we observed that the expression of the sigB gene was heterogeneous within a population of vegetative cells. Using a P sigB -SNAP Cd fusion, we showed that sigB is only expressed in a small fraction (between 10 and 20 %) of C. difficile cells during both exponential (4 h) and at the onset of the stationary growth phase (10 h) in both the wild-type and sigB mutant strains (Kint et al, in Annex). This is in agreement with the fact that C. difficile sigB expression is not autoregulated, and with the transcriptomic analysis that do not reveal differences in the expression of this gene whatever the growth stages (Saujet et al. 2011) (Kint et al, in Annex). As observed for the sigB gene, the expression of the flavodiiron protein (FDP) gene CD1157 under the control of the dual promoter σ A /σ B or only through the σ A -or σ B -dependent promoters, was also detected in a very few number of cells during stationary phase. However, no major differences were observed in the percentage of cells expressing CD1157 under the control of σ A /σ B or σ A alone. Nevertheless, the number of cells expressing CD1157 under σ B control is even lower suggesting that the expression of the CD1157 gene may be below the fluorescence detection limit. The same heterogenicity of expression was observed for genes encoding the reverse rubrerythrins (revRbrs, CD1474 and CD1524) and the second FDP (CD1623) during stationary phase, which support that the activation of σ B is heterogeneous. We noted that the expression of these genes was detected in a higher percentage of cells than the expression of sigB itself. Because these genes are only under the control of σ B , it is possible that the level of expression of the P sigB -SNAP Cd fusion may be below the fluorescence detection limit. Finally, it would be interesting to verify if the cells expressing sigB show the same σ B activity by using different reporter genes.

Additionally, the analysis of the expression of other σ B -controlled genes at the single cell level will be important to confirm the heterogeneity of σ B activity.

Such heterogeneity of σ B expression and activity may be a strategy to increase the chances of part of the population to survive when the environmental conditions become adverse (Eldar andElowitz 2010, Guldimann et al. 2017). Heterogeneity of gene expression during stationary phase was recently showed for the C. difficile toxins, where only a fraction of the population expresses tcdA (Ransom et al. 2018b). This suggests that within a bacterial population, subpopulations could be responsible for different physiological functions through "bet-hedging" strategies in order to maximize the chances of survival in hostile conditions.

Each C. difficile sub-population would invest in functions such as survival, toxin production or dissemination (Figure 59). This mechanism of "bet-hedging" might be important in C.

difficile for an efficient colonization of the intestinal tract (Kint et al. 2017). However, the

III. O 2 tolerance and ROS protection in C. difficile

III.1. Protective role of the C. difficile revRbrs and FDPs

Previous results demonstrated an increased expression of genes encoding revRbrs and FDPs when C. difficile is exposed to air [START_REF] Emerson | Microarray analysis of the transcriptional responses of Clostridium difficile to environmental and antibiotic stress[END_REF]. Surprisingly, in recent transcriptomic studies of C. difficile grown under microaerophillic conditions, these genes were not among the subset of upregulated genes (Giordano et al. 2018a[START_REF] Neumann-Schaal | Tracking gene expression and oxidative damage of O2stressed Clostridioides difficile by a multi-omics approach[END_REF] exposure and funtion as NADH oxidases [START_REF] Kawasaki | Identification of O2-induced peptides in an obligatory anaerobe, Clostridium acetobutylicum[END_REF][START_REF] Kawasaki | Adaptive responses to oxygen stress in obligatory anaerobes Clostridium acetobutylicum and Clostridium aminovalericum[END_REF].

However, the oxidase specific activities of FprA1/ FprA2 are at least 100-fold higher than that of the revRbrs (Hillmann et al. 2009b, Riebe et al. 2009). In C. difficile, the NADH oxidase activities of the FDP and the revRbrs were almost similar (Table 3). Additionally, using the same artificial system described above (NROR and Rd from E. coli), we showed that the revRbrs CD1474 and CD1524 also have NADH peroxidase activity. As observed in C. possibility that could be tested is whether the NROR and the Rd domains of the FDP CD1623 are able to reduce the revRbrs and/or the FDP CD1157.

In a previous study, FDP CD1623 showed an O 2 -reductase activity (Folgosa et al. 2018b). The O 2 reductase activity of this enzyme was higher than the observed for the revRbrs and FDP CD1157 (Table 3). However, while we observed a decreased tolerance of the ∆CD1474-∆CD1524 double mutant and the CD1157 mutant to 0.4 % O 2 , only a small difference was observed for the ∆CD1623 mutant when compared to the wild-type. At 0.4 % O 2 , the ∆CD1623-CD1157 double mutant was only slightly different from the CD1157 mutant. Future experiments at lower and higher percentages of O 2 will be performed in order to test more precisely the differences in O 2 tolerance between the mutants. When exposed to air, we observed a slight increased sensitivity of the ∆CD1623 mutant, while no differences were observed for the other mutants. The reason why the ∆CD1474-∆CD1524 double mutant and the CD1157 mutant are more sensitive to low O 2 tension but not to air is not yet clear.

The determination of the K m of each enzyme will give additional information that might help to explain these differences.

The CD1623 protein contains all the domains necessary to oxidize the NADH and to directly reduce the substrate. By contrast, it is noteworthy that an artificial system of electron transfer was used to determine the revRbrs and CD1157 activities and can lead to an underestimation of the physiological activities. In addition, only half of the CD1157 purified protein was actually active due to an incomplete association with iron and FMN that are important cofactors for the enzymatic activity. Based on these data, we cannot exclude the possibility that the O 2 -reductase activity determined for our proteins might be underestimated.

III.2. Protection against H 2 O 2

Despite the different attempts and methods we tried to analyse the role of the revRbrs and the FDPs in H 2 O 2 protection in C. difficile, we were unable to observe differences between the various mutants and the wild-type strain when exposed to this toxic compound.

Therefore, even if an H 2 O 2 -reductase activity was observed for the two revRbrs and for the FDP CD1623 (Folgosa et al. 2018b), the absence of a phenotype for the corresponding mutants suggests functional redundancy. We showed that the C. difficile sigB mutant is more sensitive to H 2 O 2 (Kint et al. 2017). In addition to the revRbrs and FDP CD1623, σ B positively controls additional genes encoding proteins such as the glutamate dehydrogenase

GluD [START_REF] Girinathan | Clostridium difficile glutamate dehydrogenase is a secreted enzyme that confers resistance to H2O2[END_REF], and thioredoxins involved in thiol homeostasis (Kint et al. 2017). These proteins, together with the revRbrs and the FDP CD1623 might contribute to the C. difficile resistance against H 2 O 2 . Moreover, C. difficile genome encodes two classical Rbrs:

CD2845 and CD0825. CD2845 is expressed under the control of the sporulation-specific sigma factor σ G and is likely not expressed during vegetative growth (Saujet et al. 2013).

However, CD0825 gene is part of an operon containing genes encoding the PerR regulator, a desulfoferrodoxin and a glutamate synthetase. A σ A -dependent promoter is present upstream of the CD0825 gene, but the control of the expression of this operon by PerR remains to be determined. A peroxidase activity of the classical Rbr from C. difficile, as well as the presence of other proteins able to scavenge H 2 O 2, might still protect C. difficile against these compound, even in the absence of the revRbrs. To confirm these hypothesis, we plan to analyse in collaboration with Prof. Miguel Teixeira's group (ITQB, Portugal) the NADH oxidase and peroxidase activities of the crude extracts of the different single and multimutants, as it has been performed in C. acetobutylicum [START_REF] Hillmann | PerR acts as a switch for oxygen tolerance in the strict anaerobe Clostridium acetobutylicum[END_REF]. We can speculate that if the peroxidase activity of crude extracts of the mutant inactivated for the two revRbrs is similar to the one observed in the wild-type strain, it would suggest that other proteins can have such activity in addition to that of the revRbrs. Additionally, this experiment might allow us to better compare the activity of the different revRbrs and FDPs enzymes without using artificial electron partners.

III.3. O 2 and ROS response in B. subtilis and clostridia

In C. acetobutilycum, σ B is absent, and PerR is involved in H 2 O 2 and O 2 resistance through the regulation of the expression of the genes encoding the revRbrs, among others [START_REF] Hillmann | PerR acts as a switch for oxygen tolerance in the strict anaerobe Clostridium acetobutylicum[END_REF]. In this bacterium, the inactivation of perR leads to prolonged aerotolerance and higher resistance to H 2 O 2 [START_REF] Hillmann | PerR acts as a switch for oxygen tolerance in the strict anaerobe Clostridium acetobutylicum[END_REF]). An increased tolerance to H 2 O 2 exposure was also observed in a strain overexpressing the genes encoding the revRbrs [START_REF] Riebe | Pathway for H2O2 and O2 detoxification in Clostridium acetobutylicum[END_REF]. Interestingly, the authors were not able to inactivate the genes encoding the revRbrs suggesting that these genes are essential and required for the survival of C.

acetobutylicum [START_REF] Hillmann | The rubrerythrin-like protein Hsp21 of Clostridium acetobutylicum is a general stress protein[END_REF]). Since we did not observe difference in the protection agaisnt H 2 O 2 between the C. difficile wild-type strain and both the double ∆CD1474-∆CD1524 and single CD1623 mutant, it would be interesting to test H 2 O 2 sensitivity using the complemented strains where we expect an overexpression of each gene through a multicopy plasmid. In B. subtilis, the oxidative stress resistance can be mediated by catalases controlled

III.4. Protection against NO

Some FDPs have been shown to scavenge nitric oxide (NO) [START_REF] Silaghi-Dumitrescu | A flavodiiron protein and high molecular weight rubredoxin from Moorella thermoacetica with nitric oxide reductase activity[END_REF][START_REF] Silaghi-Dumitrescu | A flavo-diiron protein from Desulfovibrio vulgaris with oxidase and nitric oxide reductase activities. Evidence for an in vivo nitric oxide scavenging function[END_REF], Folgosa et al. 2018a). CD1157 belongs to the class A FDP. These FDPs typically have a clear preference for O 2 [START_REF] Seedorf | F420H2 oxidase (FprA) from Methanobrevibacter arboriphilus, a coenzyme F420-dependent enzyme involved in O2 detoxification[END_REF][START_REF] Smutna | Flavodiiron protein from Trichomonas vaginalis hydrogenosomes: the terminal oxygen reductase[END_REF][START_REF] Vicente | A detoxifying oxygen reductase in the anaerobic protozoan Entamoeba histolytica[END_REF], or a dual reductase activity for NO and O 2 , as observed for example for the FprA2 from C. acetobutylicum [START_REF] Silaghi-Dumitrescu | A flavodiiron protein and high molecular weight rubredoxin from Moorella thermoacetica with nitric oxide reductase activity[END_REF][START_REF] Rodrigues | Desulfovibrio gigas flavodiiron protein affords protection against nitrosative stress in vivo[END_REF], Hillmann et al. 2009b). Contrary to FprA2, only O 2 -reductase activity was observed for CD1157 (Table 3). In addition, the C. difficile CD1623 of the class F FDP is clearly an O 2 -reductase and does not have NO-reductase activity (Folgosa et al. 2018b). To date, the determinants of substrate specificity of the FDPs remains unclear, although it has been shown

in Entamoeba histolytica that some amino acids in the catalytic center of the FDP can modulate its enzymatic activity [START_REF] Goncalves | Flavodiiron oxygen reductase from Entamoeba histolytica: modulation of substrate preference by tyrosine 271 and lysine 53[END_REF]. The alignment of C. difficile CD1157

with FDPs from other organisms reveals the conservation of Lys 53 and Tyr 263 that are characteristic of FDPs with a preference for O 2 as substrate (Figure 60). The presence of a Lys 55 and a His 267 in CD1623 suggested a specificity of this enzyme to O 2 (Figure 60). This is in agreement with the enzymatic activities found for these two enzymes mainly as O 2reductases. Regarding the two revRbrs, they did not show NO-reductase activity. So far, no report of Rbrs with NO-reductase activity has been documented. The sensitivity of our different revRbrs and FDPs mutants to NO-donor compounds was analysed but no differences were observed when compared to the wild-type strain. This result was not surprising since none of the proteins seems able to efficiently use NO as substrate. Among genes positively controlled by σ B in C. difficile, some encode proteins that may intervene in NO detoxification pathways. These proteins include a hydroxylamine reductase (CD2168) and two nitroreductases (CD1125 and CD0837). A consensus sequence recognized by σ B was found in Altogether, our results clearly demonstrate that the two revRbrs and the two FDPs play an important role scavenging O 2 . Additionally, the two revRbrs are probably part of a subset of proteins that confers resistance to oxidative stress by scavenging H 2 O 2 . Further studies will be needed to address additional candidates that combined with these proteins, constitute the reductive scavenging system for O 2 and ROS in C. difficile. Figure 61 summarizes the proposed ROS and O 2 detoxification pathways involving revRbrs, FDPs and yet non-identified proteins. Candidates for ORs might be CD0176, CD2540 and CD2709. In addition, we propose that the desulfoferrodoxin CD0827, is a candidate for the reduction of the superoxide anion in C. difficile.

III.5. Importance of O 2 , ROS and NO detoxification pathways during colonization

The inactivation of genes encoding proteins playing oxidative protective roles, results in a decrease in the virulence or colonization of the host. Candida albicans cells lacking a superoxide dismutase (SOD) are more susceptible to macrophage attack and had atenuated virulence in mice [START_REF] Hwang | Copper-and zinccontaining superoxide dismutase (Cu/ZnSOD) is required for the protection of Candida albicans against oxidative stresses and the expression of its full virulence[END_REF]. In Streptococcus agalactiae, SodA contributes to the virulence of the bacterium [START_REF] Poyart | Contribution of Mn-cofactored superoxide dismutase (SodA) to the virulence of Streptococcus agalactiae[END_REF]. Despite the important role played by FDPs and Rbrs in scavenging O 2 , H 2 O 2 and NO, studies showing their specific roles in vivo colonization are still limited. Mydel et al showed that the Rbr of Porphyromonas gingivalis ensures the proliferation of this pathogen in mice. However, the in vivo protection afforded by the Rbr was not associated with the oxidative burst responses but rather with protection agaist reactive nitrogen species (RNS) [START_REF] Mydel | Roles of the host oxidative immune response and bacterial antioxidant rubrerythrin during Porphyromonas gingivalis infection[END_REF]). In C. difficile, the inactivation of sigB results in a major colonization defect. However, σ B is a master regulator controlling a lot of different processes, including O 2 tolerance, ROS and NO protection (Kint et al. 2017). The specific role of the C. difficile revRbrs and the FDPs in colonization is a crucial point that will be addressed in future work. However, when all the alternative survival strategies fail, the last resource of this bacterium is to differentiate into a highly resistant dormant cell type, the endospore [START_REF] Stephens | Bacterial sporulation: a question of commitment?[END_REF]. In C.

difficile, sporulation is negatively controlled by σ B . The expression of more than 200 genes involved in sporulation are derepressed in a sigB mutant and this mutant sporulates more efficiently and/or earlier than the wild-type strain (Kint et al. 2017). In B. subtilis, the overexpression of sigB results in a decreased sporulation efficiency (Reder et al. 2012a, Reder et al. 2012b). Similarly, the inactivation of rsbW or rsbX, causing σ B over-activation, inhibits the ability of B. subtilis to sporulate (Kalman et al. 1990, Benson andHaldenwang 1992). In this bacterium, σ B seems to act at the initiation of the sporulation process by influencing the phosphorylation level of the master regulator of sporulation, Spo0A. Indeed, the spo0E gene, encoding a phosphatase involved in the dephosphorylation of Spo0A~P, has a promoter recognized by σ B (Reder et al. 2012a). In C. difficile, σ B negatively controls the expression of the CD1579 gene, encoding a kinase of Spo0A. Additionally, σ B positively controls the expression of the CD1492 gene, encoding a phosphotransfer protein that seems to have a negative effect on sporulation since its inactivation increases spore production [START_REF] Childress | The Phosphotransfer Protein CD1492 Represses Sporulation Initiation in Clostridium difficile[END_REF]. These data support the idea that σ B might lead to a decrease in Spo0A~P concentration resulting in the limitation of sporulation (Figure 62). Thus, to face stressful environments encountered during infection, C. difficile may use two strategies: 1) a rapid adaptive response to stress stimuli mediated by σ B or 2) an irreversible and slower differentiation process leading to the formation of spores.

IV.2. sigB expression in sporulating cells

Surprisingly, we showed that even if σ B represses sporulation, the sigB gene continues to be expressed during this developmental process. More importantly, σ B is still active during sporulation since the genes encoding the two revRbrs, CD1474 and CD1524, the FDP CD1623, and the OR CD0176 are also expressed and proteins produced. We demonstrated actually that when the cells are grown in a medium that promotes sporulation, more than half of the vegetative cells express sigB and its targets genes (the revRbrs and the FDP CD1623)

after 24 h of growth. Although these genes are expressed in both MC and FS compartments Discussion 159 from early sporulation steps until engulfment completion, both sigB and its targets will only be expressed in the MC compartment at the end of sporulation process. The mechanism behind this differential control of expression remains unknown. However, we noted that the expression of these genes during sporulation is still dependent on sigB and, that the sigB operon including rsbV and rsbW is transcriptionally regulated by σ A and expressed during sporulation. Several studies in B. subtilis demonstrated that σ A continued to be active during sporulation in both compartments [START_REF] Fujita | Temporal and selective association of multiple sigma factors with RNA polymerase during sporulation in Bacillus subtilis[END_REF][START_REF] Li | Development of a two-part transcription probe to determine the completeness of temporal and spatial compartmentalization of gene expression during bacterial development[END_REF][START_REF] Fukushima | Transcriptional, functional and cytochemical analyses of the veg gene in Bacillus subtilis[END_REF][START_REF] Ohashi | Far different levels of gene expression provided by an oriented cloning system in Bacillus subtilis and Escherichia coli[END_REF]. Certain classes of housekeeping genes such as the sacB gene transcribed from a σ A -dependent promoter are expressed in the FS and in the MC both before and after engulfment [START_REF] Li | Development of a two-part transcription probe to determine the completeness of temporal and spatial compartmentalization of gene expression during bacterial development[END_REF]. However, expression of σ A -dependent motility genes or other genes involved in the synthesis of histidine, phenylalanine and serine were severely reduced after initiation of sporulation. These suggest that the expression of particular σ Adependent genes is subjected to additional regulatory mechanisms [START_REF] Steil | Genome-wide analysis of temporally regulated and compartment-specific gene expression in sporulating cells of Bacillus subtilis[END_REF].

Unfortunately, as far as I know, the expression of B. subtilis sigB gene has not been monitored during sporulation using fluorescence microscopy. However, based on our results

we may hypothesize that in C. difficile additional regulators might be involved in the compartmentalization of σ A -dependent gene expression. It is also possible that an unknown regulatory mechanism is involved in the control of sigB expression during sporulation.

However, we cannot exclude other possibilities such as the binding of SASPs proteins to the FS DNA at the end of sporulation that might hinder the expression of certain genes in the FS, such as the sigB operon. The analysis of the expression of C. difficile housekeeping genes and genes under the dual control of σ A and σ B , such as CD1157 encoding the FDP, during the different morphogenetic stages of sporulation will be an interesting point to address in the future. This would give new insights about the role of both σ A and σ B in sporulation.

Another interesting point to be discussed is the detection of the anti-sigma factor RsbV in the total proteome of the spore (Lawley et al. 2009b). Neither σ B nor the anti-anti-sigma factor RsbW was detected. First, this fact might suggest an increased stability of RsbV over the others. Second, the undetected σ B is in agreement with the absence of σ B activity in the FS compartment after engulfment completion.

We also demonstrated that even if sigB was expressed in more than half of the cells after engulfment completion, the expression of the gene encoding the OR CD0176 was detected only in a few cells at the same stage. Since the expression of CD0176 is also strictly dependent on σ B , this data suggests that a targeted activity of σ B might exist, associated to the Spores are resistant to O 2 but also to H 2 O 2 , hypochlorite and other disinfectants used in hospital (Fawley et al. 2007, Otter and[START_REF] Otter | Survival of nosocomial bacteria and spores on surfaces and inactivation by hydrogen peroxide vapor[END_REF]. Several factors contribute to the protection of the spore against oxidizing agents, such as the DNA saturation by SASPs, the low permeability of the spore inner membrane and, the spore coat layer [START_REF] Setlow | Spore Resistance Properties[END_REF]. The protective role of the spore coat is very likely related both with its barrier effect and with the presence of proteins able to detoxify oxidizing compounds before they reach more internal targets [START_REF] Riesenman | Role of the spore coat layers in Bacillus subtilis spore resistance to hydrogen peroxide, artificial UV-C, UV-B, and solar UV radiation[END_REF]. In B. subtilis, CotA protects spores from H 2 O 2 [START_REF] Hullo | CotA of Bacillus subtilis is a copper-dependent laccase[END_REF]). However, a protective role against ROS by SodA and the catalases CotJC and YjqC present at the spore surface has not been shown [START_REF] Henriques | Characterization of cotJ, a sigma E-controlled operon affecting the polypeptide composition of the coat of Bacillus subtilis spores[END_REF][START_REF] Henriques | Involvement of superoxide dismutase in spore coat assembly in Bacillus subtilis[END_REF]. In C. difficile, catalase activity was observed for the recombinant catalases, CotD and CotG but not for CotCB. The cotD mutant spores showed a reduction in catalase activity compared to the wild-type strain (Permpoonpattana et al. 2013). The expression of the genes encoding the revRbrs and the FDP CD1623 in the MC compartment is in agreement with the localization of the corresponding proteins at the outermost layers of the spore. These proteins could contribute to the resistance of the spore to O 2 and ROS. Indeed, the revRbrs have O 2and H 2 O 2 -reductase activities in vitro. However, contrary to the catalases that can perform their enzymatic reactions without needing other partners, the revRbrs need an electron transfer pathway and an electron donor, such as NADH. The FDP CD1623, which also has O 2 -and H 2 O 2 -reductase activities, contains all the domains and can play its enzymatic role without partners, but still needs an electron donor. It should be noted that spores being metabolically inactive produce minimal if any levels of ATP and NADH [START_REF] Korza | Levels of L-malate and other low molecular weight metabolites in spores of Bacillus species and Clostridium difficile[END_REF], Setlow et al. 2017). Whether these proteins are active and play a protective role at the spore surface is still speculative and will be investigated. Therefore, it would be interesting to compare the peroxidase activity of the spores of the different revRbrs and FDPs mutants as well their resistance to H 2 O 2 . It is also important to extend this study to other σ B targets, detected or not in the proteome of the spore outermost layers. For instance, one of the two C. reductase-CD1691 is transcribed by σ B . The thioredoxin reductase was detected in the proteome of the spore, but not to that of the spore surface layers (Lawley et al. 2009b, Abhyankar et al. 2013, Diaz-Gonzalez et al. 2015). The study of the expression of these genes and the localization of the encoding proteins might give clues about the role of σ B and its targets in spore protection.

IV.3.b. Germination and Outgrowth

The expression of the genes encoding the two revRbrs, the FDP CD1623 and the OR CD0176 in the FS compartment during sporulation might suggest importance of their role in the future vegetative cell, particularly during outgrowh. Clostridium novyi spores contain mRNA encoding a thioredoxin reductase, a glutaredoxin, a glutathione peroxidase and a Rd [START_REF] Bettegowda | The genome and transcriptomes of the anti-tumor agent Clostridium novyi-NT[END_REF]). Glutathione peroxidases have been implicated as major scavengers for H 2 O 2 [START_REF] Arthur | The glutathione peroxidases[END_REF]. Interestingly, transcripts encoding CD1631-SodA, two putative OR complexes with ferredoxin activity (CD2197-CD2199 and CD2427-CD2429), a NADH oxidase (CD2540) and a NADH-dependent flavin oxidoreductase (CD2709) have been found

in C. difficile germinating spores. The presence of some σ B -targets transcripts stored in the spore could also allow the spores to rapidly synthesize proteins that are required for germination and outgrowth in premissive oxidative environments. It would be interesting to monitor the expression of these genes during outgrowth from purified spores carrying the transcriptional SNAP fusions. In future work, the analysis of the germination efficiency of the different mutants will be also carried out in conditions mimicking the gut environment.

Another interesting possibility to be addressed is whether the localization of the revRbrs, FDPs and the OR at the outermost layers of the spore are important during outgrowth as well.

During germination and outgrowth, the spore coat layer is discarded to allow emergence of the vegetative form [START_REF] Hamilton | Comparison of the germination and outgrowth of spores of Bacillus cereus and Bacillus polymyxa[END_REF]Stubbs 1967, Abee et al. 2011). Because the revRbrs, FDPs and the OR proteins are part of the outermost layer, they might provide an arsenal that protects the cell during outgrowth, through the detoxification of toxic compounds in the surrounding environment.

IV.3.c. Mother cell protection

Recent results have raised another hypothesis for the role of σ B during sporulation.

Indeed, after 7 days of growth in sporulation medium, we observed that the double mutant inactivated for the two revRbrs produce less spores than the wild-type strain (Figure 63).

Additionally, phase-contrast microscopy analysis revealed a mixture of immature phase-dark Discussion 162 and mature spores phase-bright spores for the double mutant purified spores, while mainly phase bright spores were observed in the wild-type strain. The reason for the high amount of phase-dark spores in the ∆CD1474-∆CD1524 double mutant is not yet known. However, given the decreased tolerance of the double mutant to low O 2 tension and probably to other stresses, we hypothesize that the mutant cells might be more sensitive and more susceptible to lysis. During sporulation, the timing of the MC lysis is crucial to ensure the release of a mature spore [START_REF] Sato | Kallmann syndrome: somatic and germline mutations of the fibroblast growth factor receptor 1 gene in a mother and the son[END_REF]. A premature lysis of the sporulating cell might typically prevent for a proper formation and maturation of the spore. This might explain, in part, the decreased spore yield for the double mutant inactivated for the two revRbrs. In the colon, C.

difficile sporulates in conditions where the cells are exposed to inflammation. These proteins, not only protect the cell against O 2 and possibly H 2 O 2 during growth, but they also might play an important protective role for the endospore during sporulation. The activity of σ B and the production of the revRbrs in the MC until the last steps of spore formation might be important to protect the cell before the mature spore is formed and released to the environment. In order to address our hypothesis, we plan to monitor the sporulation of the double mutant during the 7 days in SMC plates and combine it with live/death cell microscopy experiment. In addition, we plan to test the effect of stressful compounds such as H 2 O 2 on the sporulation process of the wild-type and the double mutant. Very few studies exist concerning the role of revRbrs.

Two revRbrs of around 90% identical to the revRbrs (CD1474 and CD1524) of C. difficile, were identified in the exosporium of C. sordelli spores [START_REF] Rabi | Clostridium sordellii outer spore proteins maintain spore structural integrity and promote bacterial clearance from the gastrointestinal tract[END_REF]. However, the mechanism behind their synthesis is not yet known. The sigB operon is conserved in this bacterium, however its role has not been yet studied. Studies on the expression and role of revRbrs in other spore forming bacteria will be important to help to elucidate their role during sporulation.

V. Conclusion

This work brought new information on 1) key enzymes used by C. difficile to scavenge O 2 and H 2 O 2 and involved in oxidative stress tolerance and 2) the structure and morphogenesis of the C. difficile spore coat layers. Indeed, after spore germination in the small intestine, the vegetative cells of C. difficile will multiply and colonize the colon as a nutrient-rich environment, while being exposed to toxic compounds that can affect their growth (Figure 64). Moreover, the production and secretion of toxins by C. difficile lead to a strong inflammation process that promotes the production of antimicrobial oxidizing compounds by the host immune cells such as ROS, NO and RNS. In addition, an increasing gradient of O 2 exist from the intestinal lumen (anaerobic) to the mucosa (approximately 5% O 2 ), which can increase again and/or be disturb after an antibiotic treatment. The implementation of a stress response mediated by σ B appears as a first survival strategy. σ B , which is active in a fraction of the vegetative cells, inhibits sporulation and controls genes essential to guarantee an efficient colonization of the gut. Among them, are the revRbrs-and FDPs-encoding genes (Figure 64). In this work, we showed that these proteins are essential determinants for the ability of C. difficile to tolerate and survive in the presence of low O 2 tension. Whether these proteins play a role in vivo in the detoxification of H 2 O 2 will be addressed in the future. When the conditions become too hostile, sporulation, a long and irreversible process, is then implemented to produce spores. Interestingly, even if σ B represses sporulation, σ B -mediated stress response and sporulation might not be mutually exclusive processes. The synthesis of the revRbrs in the MC compartment, controlled by σ B during sporulation, might protect the cell against oxidizing components by inactivating them before the spore is ready to be released (Figure 64). The C. difficile spore is the highly infectious form of this bacterium and serves as means of survival during transmission from one host to another. In this work, we characterized a new spore protein, CotL that is essential for the assembly of the spore coat layers and ultimately, for the formation of a resistant spore (Figure 64). We found that CotL is located inside the coat of the C. difficile spore, and demonstrated that its role during coat assembly is to mediate the encasement of other coat components around the FS during sporulation. However important questions still remain and particularly on how CotL, together with SpoIVA and SipL, interact with all the proteins that they direct to the surface of the developing spore. Nevertheless, the present study has expanded our knowledge on C. difficile spore morphology, highlighting that the coat assembly pathway in C. difficile presents substantial differences relative to that of B. subtilis.

Introduction

Clostridium difficile, a gram-positive, anaerobic and spore-forming bacterium, is the main cause of antibiotic-associated diarrhoea and of pseudomembranous colitis, a potentially lethal disease. In the early 2000s, the incidence and severity of C. difficile infection (CDI)

have increased in North America and Europe owing to the emergence of new isolates of the BI/NAP1/027 lineage (Rupnik et al. 2009, Wiegand et al. 2012). C. difficile is acquired from the environment through the ingestion of spores: a form of resistance, transmission and dissemination of this bacterium (Deakin et al. 2012). Disruption of the normal intestinal microbiota, notably by antibiotics, induces substantial changes in metabolite pools (Theriot et al. 2014). These changes include an increased concentration of bile salts that triggers spores Hecker et al. 2007, Guldimann et al. 2016), plays a crucial role in the regulation of these different processes in C. difficile (Kint et al. 2017). Accordingly, sigB inactivation leads to a higher sensitivity to several stresses the bacterium copes within the gut such as acidic pH, antimicrobial peptides, ROS, NO, RNS and O 2 . In this enteropathogen, σ B controls the expression of about 25 % of the genes at the onset Annexes 202 of stationary phase (Kint et al. 2017). In addition, the colonisation fitness of the sigB mutant is greatly affected in dixenic mice when compared to wild-type strain (Kint et al. 2017).

Despite σ B involvement in stress protection and in gut colonization, little is known about the mechanisms involved in its activation in C. difficile.

Since the expression of σ B regulon represents a major cost for the bacteria, σ B activity is tightly controlled. In other firmicutes, σ B activity is modulated by a well-characterised posttranslational mechanism called partner switching, in which the interaction and activation of proteins are mediated by reversible threonine and serine phosphorylation (Hecker et al. 2007, de Been et al. 2011). In unstressed cells and during exponential growth phase, the anti-σ factor RsbW phosphorylates the anti-anti-σ factor RsbV blocking its activity (Alper et al. 1996). This enables RsbW to sequester σ B thereby preventing its association with RNA polymerase core enzyme. During stationary growth phase or when cells are exposed to a stress, a signal is conveyed to specific PP2C phosphatase(s) that dephosphorylate(s) RsbV.

The unphosphorylated form of RsbV interacts with RsbW, leading to the release and activation of σ B and the transcription of its target genes (Yang et al. 1996, Vijay et al. 2000, van Schaik et al. 2005). The stress signals triggering σ B activation differ among the firmicutes. They can be divided into two groups i) environmental stresses such as ethanol, elevated temperature, hyperosmolarity and low pH (Boylan et al. 1993, Voelker et al. 1995, van Schaik et al. 2004, Pane-Farre et al. 2006) ii) energetic stresses like carbon or phosphate starvation, O 2 deprivation and ATP depletion (Voelker et al. 1995, Vijay et al. 2000, Zhang and Haldenwang 2005).

In this work, we sought to elucidate how σ B becomes active in C. difficile. We first studied the regulation of sigB expression and then focused on the partner switching mechanism controlling σ B activity. We identified the PP2C phosphatase that acts upstream of this partner switching and turn σ B active. We sought to identify other actors/sensors and their putative role in the σ B activation pathway. Finally, sigB expression and σ B activity was studied at a single cell level within a population of C. difficile vegetative cells using fluorescent microscopy. Altogether the results obtained bring new insights into σ B activation signalling pathway in C. difficile and highlight the differences compared to other firmicutes. Gene function is indicated below each gene. TSS mapping experiment indicated the presence of a σ A -dependent promoter upstream of CD0007 (Soutourina et al, 2013). The -35 and -10 boxes as well as the TSS are indicated in bold. PCR were both realised on RNAs without reverse transcription (lane 1, 3, 6 and 8) and on cDNA synthesised from the same RNAs by reverse transcription using IMV389 primer (lane 2, 4, 7 and 9). PCR was then performed with primers annealing in CD0007 and CD0008 (IMV774/OS548, lane 1 and 2), in CD0008 and CD0009 (OS547/IMV777, lane 2 and 3), in CD0009 and CD0010 (IMV776/IMV779, lane 6 and 7) and in CD0010 and CD0011 (IMV778/IMV389, lane 8 and 9). Lane 5: Ladder.

Table 1: Expression of sigB operon in the sigB mutant and complemented strain.

To study σ B autoregulation, cells were harvested after 10 h of growth in TY. The results represent the mean of three independent experiments.

Results

Genetic organisation of sigB operon

The genetic environment of sigB gene is rather conserved among the firmicutes harbouring this σ factor such as Bacillus subtilis, Bacillus cereus, Listeria monocytogenes and Staphylococcus aureus (Wise and Price 1995, Wu et al. 1996, Wiedmann et al. 1998, van Schaik et al. 2004). The sigB gene forms an operon with the genes encoding the proteins involved in the control of its activation, i.e. the anti-anti-sigma factor and the anti-sigma factor, RsbV and RsbW, respectively. In C. difficile, sigB (CD0011) is the last gene of a locus containing five genes (CD0007-0011). CD0007 and CD0008 encode a 98 amino acid protein with a functional uncharacterized YtxH conserved domain (pfam12732) and an 86 amino acid protein without identified domain, respectively. Using THMM and Signal-P software, no signal peptides were detected for these small proteins, but one trans-membrane helix was predicted for CD0007 (amino acids 12 to 29) and for CD0008 (amino acids 5 to 24). CD0009 and CD0010 share 35 % and 39 % of identity with RsbV and RsbW of B. subtilis, respectively. To determine whether these five genes form an operon, we performed RT-PCR experiments. PCR products were detected using pairs of primers located in adjacent genes, with RNAs processed by reverse transcriptase but not with untreated RNAs, serving as a negative control (Fig. 1). Since, DNA fragments were amplified only if neighbouring genes were co-transcribed, these results indicate that sigB is the last gene of a five-gene operon containing rsbV and rsbW.

Absence of auto-regulation and constitutive expression of sigB

A previous genome-wide transcriptional start site (TSS) experiment mapped a single TSS in sigB operon located 34 bp upstream of the first codon of CD0007 (Soutourina et al. 2013). Upstream of this TSS, we found a -10 (TATAAT) box similar to the consensus sequence of σ A -dependent promoters while the -35 (TAGGAT) region is less conserved (Fig. 1). This promoter clearly differs from the consensus of σ B -dependent promoters recently defined in C. difficile (Kint et al. 2017). The presence of a single σ A -dependent promoter suggested that sigB expression is not auto-regulated in C. difficile, contrary to the positive control of its own transcription observed in other firmicutes. Accordingly, we did not detect any σ B -dependent change of expression of CD0007, CD0008, rsbV or rsbW by qRT-PCR (Table 1). These results are in agreement with our previous transcriptomic study (Kint et al. 2017). To determine whether expression of sigB operon varies during C. difficile growth, we of rsbV, rsbW and sigB genes and of some σ B target genes between 630∆erm + P tet -sigB and 630∆erm + pDIA6103 without aTc (black), with 25 ng.ml -1 (dark gray) or 200 ng.ml -1 of aTc (light gray). These results are the means of 2 independent experiments.

Annexes 204 monitored its expression by qRT-PCR during growth in TY. We did not observe significant difference in sigB operon expression throughout the growth (data not shown). Altogether, these results highlight an absence of σ B auto-regulation and a constitutive sigB operon expression in C. difficile.

Toxicity of sigB overexpression

As the expression of σ B regulon represents a major cost for the bacteria, sigB expression and/or σ B activity must be tightly controlled. To determine the effect of sigB overexpression, we cloned this gene into pDIA6103 plasmid allowing its expression under the control of P tet promoter, which is inducible in presence of anhydrotetracycline (aTc) (Fagan andFairweather 2011, Soutourina et al. 2013). Addition of increasing concentrations of aTc in the culture had no effect on the growth of the wild-type strain harbouring pDIA6103 empty vector (Fig. 2A). By contrast, the growth of the wild-type strain containing P tet -sigB was reduced in presence of 25 ng.ml -1 of aTc and almost abolished from 50 ng.ml -1 (Fig. 2B). To monitor the level of sigB overexpression, we extracted RNA after a 4 h growth in TY without aTc or containing either 25 ng.ml -1 or 200 ng.ml -1 . Without aTc, we did not observe difference in rsbV, rsbW or sigB gene expression between the wild-type strain containing or not P tet -sigB (Fig. 2C). By contrast, sigB gene was overexpressed 400-and 1,400-fold compared to its expression level detected in strain harbouring pDIA6103 in presence of 25 and 200 ng.ml -1 of aTc, respectively (Fig. 2C). Of note, sigB expression increased only 10fold when sigB was expressed under the control of its own promoter and the growth of this strain was not affected (Table 1 and data not shown). These results indicated that a proper expression level of sigB is important for the fitness of C. difficile. In agreement with the absence of sigB operon auto-regulation, we did not observe increased expression of rsbV and rsbW genes (Fig. 2C). Moreover, while sigB is drastically over-expressed, the expression of several C. difficile σ B -target genes transcribed from a σ B -dependent promoter only moderately increased. Indeed, the expression of CD1623 and CD1157 encoding flavodiiron proteins likely involved in the oxidative stress response (Folgosa et al. 2018b) as well as CD1474 and CD0176, encoding a reverse rubrerythrin and an NADH rubredoxin reductase, respectively, was upregulated only up to 5-fold (Fig. 2C). The different expression level between sigB and its target genes highlights a post-transcriptional mechanism involved in the control of σ B activity. Growth controls were performed with plates supplemented with NaOH or plates incubated in anaerobiosis atmosphere for NO tests and low O 2 tolerance assay, respectively. These pictures are representative of 3 independent experiments. C) Effect of rsbV deletion or rsbW overexpression on the expression of sigB operon genes and σ B target genes at the onset of stationary phase. Fold change of the differential expression of the genes belonging to the sigB operon and σ B target genes involved in oxidative and nitrosative stress management in 630∆erm ∆rsbV + pDIA6103 (black), 630∆erm ∆rsbV + P tet -rsbV (light gray) and 630∆erm + P tet -rsbW (dark gray) compared to 630∆erm + pDIA6103. These results are the mean with the standard deviation of 4 independent experiments.

σ B activation by a partner switching mechanism

According to their role in the release and activation of σ B in B. subtilis and in other firmicutes (Boylan et al. 1992, Benson andHaldenwang 1993b), we speculated that RsbV and RsbW positively and negatively control σ B activation, respectively. To test this hypothesis, we first deleted rsbV gene using allelic exchange (Ng et al. 2013, Ehsaan et al. 2016). As observed for sigB mutant (Kint et al. 2017), the growth of ∆rsbV mutant is not affected in TY (data not shown). However, we observed a higher sensitivity to diethyl-amine NoNoate (DEA/NoNoate), a NO-donor compound, and a lower tolerance to low O 2 tension for ∆rsbV mutant compared to the wild-type strain and the complemented strain (carrying P tet -rsbV) (Fig. 3A and3B). These results are reminiscent of the phenotypes previously observed in sigB mutant supporting the positive role of RsbV on σ B activity control.

Since RsbW sequesters σ B and inhibits its activity and because of the growth defect observed in the strain overexpressing sigB (Fig. 2), we assumed that rsbW deletion might be toxic in C. difficile as observed in B. subtilis (Benson and Haldenwang 1992, Boylan et al. 1992, Benson and Haldenwang 1993b). Therefore, to conclude on the role of RsbW in σ B activation, we cloned rsbW gene into pDIA6103 to induce its expression upon aTc addition.

No growth defect was observed when we overexpressed rsbW with aTc concentrations below 200 ng.ml -1 (Fig. S1). However, the strain overexpressing rsbW showed a higher sensitivity to DEA/NO compared to the wild-type strain even in presence of low aTc concentration (25 ng.ml -1 ) (Fig. 3A). Under these conditions, rsbW overexpression resulted in a lower O 2 tolerance but to a lesser extent than rsbV deletion (Fig. 3B). Altogether, these results suggest a negative role of RsbW on σ B activity control as observed in other firmicutes.

To confirm the involvement of RsbV and RsbW in σ B activation, we monitored by qRT-PCR the expression of several σ B -target genes at the onset of stationary phase, when σ B is fully active (Kint et al. 2017). The deletion of rsbV did not affect significantly the expression of sigB operon except for rsbV itself (Fig. 3C). However, the expression of CD1474, CD0176 and CD1623 encoding proteins likely involved in O 2 or ROS detoxification pathways (Kint et al. 2017, Folgosa et al. 2018b) decreased in ∆rsbV mutant compared to wild-type strain (Fig. 3C). Similarly, we observed a 2-fold overexpression of rsbW in presence of 25 ng.ml -1 of aTc, that leads to a lower expression of the different σ B -target genes but to a lesser extent than rsbV inactivation. The low level of rsbW overexpression (2-fold) might explain the reduced effect on σ B regulon expression (Fig. 3C) and the moderate phenotype observed in presence of 0.4 % of O 2 compared to ∆rsbV mutant (Fig. 3B). As rsbV deletion did not modify the expression level of sigB (Fig. 3C), we wanted to determine whether it could affect the amount of σ B protein. For this purpose, we expressed a gene encoding a HA-tagged σ B protein under the control of its own promoter in 630∆erm, sigB or ∆rsbV mutant strains. Cellular extracts were prepared from overnight cultures and σ B -HA was detected by Western blot using an antibody raised against HA tag. We detected a same amount of σ B -HA protein in the different genetic backgrounds indicating an absence of effect of rsbV deletion on the amount of σ B -HA (Fig. S2A). This suggests that RsbV acts most likely at the level of σ B activity through a post-translational mechanism. Thus, these results support that the partner switching mechanism described in other firmicutes (Yang et al. 1996) is conserved in C. difficile and enables a proper σ B activation and σ B regulon expression.

Conservation of the protein-protein interactions involved in the partner switching mechanism

The partner switching mechanism involves protein-protein interactions between RsbV, RsbW and σ B and relies on the phosphorylated state of RsbV. The amount of the anti-anti-σ factor RsbV~P is dependent on the kinase activity of the anti-σ factor RsbW as well as on the activity of specific phosphatase(s) (Dufour and Haldenwang 1994, Alper et al. 1996, Yang et al. 1996, Vijay et al. 2000). To determine the existence of interactions between RsbV, RsbW and σ B of C. difficile, we used a Bacterial Adenylate Cyclase Two Hybrid (BACTH) system, which is based on the interaction-mediated reconstruction of a cyclic AMP (cAMP) signalling cascade. Proteins of interest, which are genetically fused to two fragments (T25 and T18) of the catalytic domain of Bordetella pertussis adenylate cyclase, are co-expressed in BTH101 Escherichia coli cya strain (Karimova et al. 1998). Their interaction results in a functional complementation between T25 and T18 fragments, leading to cAMP synthesis that activates the expression of lacZ. DNA fragments encoding RsbV, RsbW and σ B were cloned into the four BACTH expression vectors to generate recombinant plasmids expressing hybrid proteins in which the examined polypeptides were fused at the N-terminus or the C-terminus of either T25 or T18 fragments (See Experimental procedures). To test the efficiency of functional complementation between the different hybrid proteins, ß-galactosidase activity was tested.

As shown in Table 2, RsbW associates with both RsbV, and σ B . By contrast, no association between RsbV and σ B have been observed in our assays (Table 2). These results indicate that proteins interactions involved in the partner switching mechanism and observed in other firmicutes are conserved in C. difficile.

Annexes 207

Identification of PP2C phosphatase CD2685

In other firmicutes, the dephosphorylation of RsbV by specific PP2C phosphatase(s) enables its interaction with RsbW and triggers the activation of σ B (Yang et al. 1996, Vijay et al. 2000, van Schaik et al. 2005)

CD2685 in σ B activation

To address the role of CD2685 in the signalling pathway leading to σ B activation, we deleted CD2685 gene by allele genetic exchange. We also complemented ∆CD2685 mutant with CD2685 gene expressed under the control of P tet promoter. The growth of ∆CD2685 mutant was not affected in TY medium (data not shown). However, we observed a growth defect in the complemented strain with 50 ng.ml -1 of aTc, and growth was almost abolished when higher concentrations were added (Fig. S1). These results were also observed in 630∆erm strain harbouring the same plasmid, suggesting that CD2685 over-expression is toxic for C. difficile (Fig. S1). Since 25 ng.ml -1 of aTc did not affect the growth of ∆CD2685 complemented strain, we decided to perform the following experiments in this condition.

To study whether CD2685 indirectly controls σ B targets expression, we extracted RNA from the wild-type strain, the ∆CD2685 mutant and the complemented strains harvested at the onset of stationary phase and we performed qRT-PCR experiments. Interestingly, although the expression of genes belonging to sigB operon was not affected, the expression of several σ B target genes including CD1474, CD0176 and CD1623 decreased 35-, 58-and 8-fold in Effect of rsbZ deletion of or its phosphatase domain (rsbZ∆pho) on the expression of sigB operon genes and σ B target genes at the onset of the stationary phase. Fold change of the differential expression of the genes belonging to sigB operon, rsbZ operon and of σ B target genes involved in oxidative and nitrosative stress management in 630∆erm ∆rsbZ + pDIA6103 (black), 630∆erm ∆rsbZ + P tet -rsbZ (light gray) and 630∆erm ∆rsbZ + P tet -rsbZ∆pho (dark gray) compared to 630∆erm + pDIA6103. Low O 2 tolerance (B) and NO stress assays (C). Serial dilutions of the different strains were spotted on TY plates containing 25 ng.ml -1 of aTc. Plates were incubated 64 h in presence of 0.4 % of O 2 (B) or 24 h at 37°C for NO tests (C). Growth controls were performed in anaerobiosis atmosphere or in presence of NaOH for low O 2 tolerance assays and NO tests, respectively. D) RsbV phosphorylation state in wild-type and ∆rsbZ backgrounds. Cellular extracts were run on 12 % acrylamide gel containing 50 µM of Phos-Tag™ compounds allowing the separation of phosphorylated and non-phosphorylated proteins. Western blot was performed with an antibody raised against the HA tag. Loading control was realized using a 12 % acrylamide gel followed by Coomassie staining. qRT-PCR results are the mean with the standard deviation of 4 independent experiments and pictures are representative of 4 independent experiments.

Annexes

208

∆CD2685 mutant compared to the wild-type strain (Fig. 4A). The expression of genes encoding proteins possibly involved in nitrosative stress protection such as the second flavodiiron protein (CD1157) and the nitro-reductase (CD1125), also decreased when CD2685 was deleted (Fig. 4A). These results are in agreement with the lower O 2 tolerance (Fig. 4B) and with the higher sensitivity to DEA/NO (Fig. 4C) of ∆CD2685 mutant compared to the wild-type and the complemented strains.

To test whether CD2685 deletion can impact σ B in a post-transcriptional way, we expressed a gene encoding a σ B HA-tagged protein in ∆CD2685 mutant. No difference in σ B -HA production in stationary phase was detected between the wild-type and ∆CD2685 mutant strains (Fig. S2A lane 2 and 5). Altogether, these results show the indirect involvement of CD2685 on the control of σ B regulon expression. Based on these results, we renamed the CD2685 phosphatase RsbZ for regulation of σ B protein Z.

RsbZ controls σ B activity by dephosphorylating RsbV

According to its PP2C phosphatase domain, we assumed RsbZ is involved in σ B activation by dephosphorylating the anti-anti sigma factor RsbV. To confirm the interaction between these two proteins, we used the BACTH system described above. No association have been detected between RsbZ and RsbW or RsbZ and σ B . By contrast, a high βgalactosidase activity was quantified from the hybrids obtained with RsbZ and RsbV supporting RsbZ is able to associate with RsbV (Table 3). This result highlighted the possible role of RsbV~P as a RsbZ substrate. To study the phosphorylation state of RsbV, we expressed a gene encoding a RsbV HA-tagged protein under the control of P tet promoter in both 630∆erm and ∆rsbZ genetic backgrounds. Cellular extracts prepared from both strains after 4 h of growth in presence of 100 ng.ml -1 of aTc were separated with the Phos-Tag™ compound that allows the discrimination of phosphorylated and unphosphorylated proteins.

By Western blot analysis using an antibody raised against the HA tag, we only detected the phosphorylated form of RsbV (RsbV~P) in ∆rsbZ background while we observed both an intense signal corresponding to the unphosphorylated form of RsbV and a faint signal corresponding to RsbV~P in 630∆erm background (Fig. 4D). These results suggest that RsbZ is responsible for the dephosphorylation of RsbV~P in C. difficile.

To confirm the requirement of the PP2C phosphatase domain for RsbZ activity and for σ B activation, we expressed rsbZ gene that lacks the 3' part of the gene corresponding to the PP2C phosphatase domain (rsbZ∆pho) under the control of P tet promoter in mutant. The growth of ∆rsbZ expressing rsbZ∆pho gene was not or slightly affected by the addition of Annexes 209 increasing amounts of aTc (Fig. S1). We confirmed by Western blot that RsbZ∆pho HAtagged protein was produced using an antibody raised against the HA tag (Fig. S2B). We then compared the ability of both the truncated RsbZ and the full-length RsbZ proteins to complement ∆rsbZ mutant. In presence of 25 ng.ml -1 aTc, rsbZ was expressed at a wild-type level in both strains (Fig. 4A). While complementation of ∆rsbZ mutant with a gene encoding the full-length protein restored σ B target gene expression at a wild-type level, complementation of this mutant with a gene encoding the truncated RsbZ∆pho form did not (Fig. 4A). These results were in agreement with the higher DEA/NO sensitivity and the lower O 2 tolerance observed in ∆rsbZ expressing only rsbZ∆pho (Fig. 4B and 4C lane 4).

Altogether, these results confirm that RsbZ phosphatase activity is essential for σ B activation.

Regulation of rsbZ expression and identification of the transcriptional start site

In C. difficile, a tight control of sigB expression and of σ B activation is needed to allow a proper growth of the cells. Since RsbZ is involved in σ B activation by RsbV dephosphorylation and because rsbZ overexpression leads to a growth defect (Fig. 4D and S2), we hypothesised that rsbZ expression could be regulated according to the growth phase.

Therefore, we monitored rsbZ expression at different time point during C. difficile growth.

The rsbZ expression level did not differ according to the growth phase suggesting a constitutive expression (data not shown) as observed for rsbP in B. subtilis (Vijay 2000). The rsbZ gene belongs to a locus containing also CD2684 and CD2686 (Fig. S3). To identify rsbZ TSS and to determine if CD2684 and rsbZ form an operon, we performed a 5' RACE experiment (see Experimental procedures). When we synthesized a cDNA using a primer annealing in CD2684 located downstream of rsbZ (NK134, Fig. S3), we identified a single TSS located 30 bases upstream of rsbZ start codon. Thus, we inferred that these two genes are co-transcribed and form an operon. We deduced -10 (TATGTT) and -35 (TTGTTT) boxes sharing similarities with the consensus sequence recognised by σ A . Previous studies showed that expression of rsbZ and CD2684 is positively controlled by the first sporulation sigma factor, σ F (Fimlaid et al. 2013). Indeed, sigF inactivation leads to a 3-and 5-fold decreased expression of rsbZ and CD2684, respectively (Fimlaid et al. 2013). Since CD2686 located upstream of rsbZ is transcribed from a σ F -dependent promoter (Saujet et al. 2013), the control of rsbZ operon expression by σ F is likely due to a transcription read-through from CD2686 promoter (Fig. S3). To test whether the control of rsbZ operon by σ F has an impact on O 2 tolerance, we exposed sigF mutant to 0.4 % of O 2 . We did not observe decrease in O 2 tolerance for sigF mutant compared to wild-type strain in our conditions (data not shown) (Fagan and Fairweather 2011) was inserted into the chromosome between rsbZ and CD2684 allowing CD2684 expression of under the control of P tet and its induction following aTc addition. B) CD2684 is essential in C. difficile. Growth phenotype of 630∆erm (left part) or sigB mutant (right part) carrying a chromosomal P tet CD2684 copy on TY plates without aTc or with 10 and 25 ng.ml -1 of aTc after 24 h at 37°C. C) Growth of 630∆erm chromosomal P tet CD2684 in TY with different aTc concentrations. D) Effect of CD2684 expression shut down on the expression of sigB operon and σ B target genes. Fold change of the differential expression of sigB, of genes of rsbZ operon and of different σ B target genes encoding proteins involved in oxidative and nitrosative stress management between 630∆erm chromosomal P tet CD2684 grown without aTc or with 10 ng.ml -1 of aTc. The graph shows the expression fold change after 4 h (black) or 10 h (light gray) of growth. Pictures are representative of 3 independent experiments and growth results represent the mean of 3 independent experiments. qRT-PCR results are the mean with the standard deviation of 3 independent experiments.

suggesting that the level of rsbZ expression from its own promoter is sufficient to trigger σ B activation.

Study of the role of CD0007 and CD0008

In other firmicutes, genes present in sigB operon are often involved in σ B activation by acting upstream of the PP2C phosphatase (Kang et al. 1996, de Been et al. 2010, Martinez et al. 2010). Since CD0007 and CD0008 belong to sigB operon (Fig. 1), we studied the possible role of these two proteins in σ B activation in C. difficile. We constructed both ∆CD0007 and ∆CD0008 single mutants as well as ∆CD0007-CD0008 double mutant and exposed these strains to low O 2 tension and to NO donor compounds. No difference in growth or in NO or O 2 sensitivity was observed between wild-type strain and the 3 mutants (Figure S4A andB).

These results suggest that neither CD0007 nor CD0008 plays a positive role in σ B activation in the tested conditions. In agreement, the expression of the σ B -target genes was not or only slightly affected by the deletion of CD0007 and/or CD0008 at the onset of stationary phase (Fig. S4C). Based on these results, we conclude that CD0007 and CD0008 are not involved in σ B signalling pathway in C. difficile.

CD2684 encodes an essential iron-sulphur flavoprotein

Proteins whose genes cluster with the gene(s) encoding the phosphatase(s) involved in RsbV dephosphorylation have also been shown to take part in the control of σ B activity in other firmicutes (Brody et al. 2001, Chen et al. 2012). Thus, we assumed the iron-sulphur flavoprotein, CD2684, could contribute to σ B activation in C. difficile. Despite several attempts, we were unable to inactivate CD2684 by allele exchange or ClosTron (Heap et al. 2007, Heap et al. 2010) suggesting that CD2684 could be essential in C. difficile. Thus, using the allele exchange strategy, we integrated into the chromosome of both 630∆erm and sigB mutant strains the entire P tet element from pRPF185 upstream of the translational initiation codon of CD2684 (Fig. 5A). This insertion enabled us to control CD2684 expression according to the amount of aTc added. Both 630∆erm and sigB mutant strains carrying P tet -CD2684 were unable to grow on plates in the absence of aTc (Fig. 5B). While 10 ng.ml -1 of aTc enabled growth restoration in the wild-type strain, 25 ng.ml -1 of the inducer was required to observe a full restoration of growth in sigB mutant (Fig. 5B). Based on these results, we conclude that CD2684 is an essential gene in C. difficile independently of the presence of σ B .

In liquid culture, 630∆erm strain harbouring a chromosomal copy of P tet promoter upstream of CD2684 showed a growth defect in the absence of aTc after 6 h (Fig. 5C). After 10 h, the Annexes 211 final OD 600nm value was lower for cultures grown in absence than in presence of the inducer (Fig. 5C). We then monitored by qRT-PCR the level of CD2684 expression in wild-type strain at two different time points (4 h and 10 h) with or without 10 ng.ml -1 of aTc. As observed in Fig. 5D, the expression level of CD2684 decreased by a 66-fold factor after 4 h of growth in absence of aTc. These results confirmed a shutdown of CD2684 expression in absence of aTc even if we failed to detect an impact on growth at 4 h. We assumed that the observed residual growth without aTc resulted from the stability of CD2684 protein produced during the overnight liquid culture done in presence of aTc. In addition, CD2684 expression decreased only 4-fold after 10 h for cultures grown without aTc (Fig. 5D). This lower ratio for CD2684 expression after 10 h of growth might be due to aTc degradation. At 10 h, a depletion of the amount of CD2684 protein might be responsible for the growth arrest observed.

CD2684 depletion prevents σ B activation

To study whether CD2684 is involved in the control of σ B activity, we compared the expression level of sigB, rsbZ and some σ B target genes between the strain harbouring a chromosomal copy of P tet upstream of CD2684 grown with or without 10 ng.ml -1 of aTc. We extracted RNAs after 4 h and 10 h of growth, i.e. when the growth of the strain was not (4 h) or was affected (10 h) by the absence of aTc. At 4 h, apart from CD2684 expression shutdown, we did not observe any significant difference change in the expression level of the tested genes (Fig. 5D). After 10 h of growth in absence of aTc, sigB expression was not affected while rsbZ expression slightly decreased with a 3-fold factor. Strikingly, we observed a more pronounced decreased expression of σ B target genes in absence of aTc (Fig. 5D). Indeed, expression of both CD1623 and CD0176 decreased 25-fold while expression of CD1474, CD1157 and CD1125 decreased 8-, 9-and 7-fold, respectively (Fig. 5D). These results strongly suggest that CD2684 depletion prevents σ B activation in C. difficile at the onset of stationay phase and support the involvement of this protein in σ B signalling pathway.

Heterogeneity of sigB expression and σ B activity in a population of C. difficile cells

To follow sigB operon expression at a single cell level within C. difficile population during both exponential and stationary phases, we fused the promoter region located upstream of CD0007 to the SNAP Cd gene (Pereira et al. 2013, Cassona et al. 2016). To monitor σ B activity, we also fused the promoter region of CD1474 that harbours a σ B consensus sequence (Soutourina et al. 2013, Kint et al. 2017) to SNAP Cd gene. Expression of P sigB -SNAP Cd and Figure 6: P sigB -SNAP Cd and P CD1474 -SNAP Cd expression during the growth. Quantitative analysis of fluorescence intensity in arbitrary units (A.U.) in vegetative cells expressing P sigB -SNAP Cd (A and B) or P CD1474 -SNAP Cd (C and D) in 630∆erm strain (red) and sigB mutant (black) after 4 h and 10 h of growth. 100 cells were scored at each time point for each strain and the numbers represent the average of fluorescence intensity. The schemes below the graphs represent the number of cells showing no (pixel values < 9 A.U.), weak (between 9 and 19 A.U.) or strong (>19 A.U.) expression of the reporter fusion. The pictures on the right side represent a field of observed cells from both the 630Δerm strain and the sigB mutant. AF: auto-fluorescence. These data are representative of 3 independent experiments. Scale bar, 1 µm. P CD1474 -SNAP Cd fusions were monitored in vegetative cells of both 630∆erm and sigB mutant strains by fluorescence microscopy using the SNAP substrate TMR-Star. In agreement with the σ A promoter found upstream of CD0007, P sigB -SNAP Cd expression was detected in both genetic backgrounds without significant difference (Fig. 6A and6B). Strikingly, P CD1474 -SNAP Cd expression was already detected in 630∆erm background after 4 h of growth (Fig. 6C). Since P CD1474 -SNAP Cd expression was completely abolished in sigB mutant, these results confirmed that σ B is strictly required for CD1474 expression (Kint et al. 2017) and indicated that σ B is active in C. difficile from the exponential growth phase. While P sigB -SNAP Cd expression was rather constant between 4 h and 10 h (Fig. 6A and6B), P CD1474 -SNAP Cd expression was induced at the onset of the stationary phase as observed by the increased average fluorescence intensity (Fig. 6D). This result confirms induction of σ B activity at the onset of the stationary phase in C. difficile, as observed in other firmicutes.

Interestingly, both P sigB -SNAP Cd and P CD1474 -SNAP Cd expressions were heterogeneous within C. difficile. Indeed, P sigB -SNAP Cd expression was detected in both backgrounds only in a small fraction of vegetative cells at the two tested time points (between 10 and 20 % when weak and strong expression were added) (Fig. 6A and6B). Similarly, we observed that during the exponential growth phase, 45 % of cells carrying the P CD1474 -SNAP Cd fusion in the 630∆erm background had a weak and strong expression level (Fig. 6C). By contrast, the percentage of cells that displayed P CD1474 -SNAP Cd expression increased at the onset of stationary phase to reach 75 % (Fig. 6D). In addition, the cells fraction with a strong signal was 3-fold higher at 10 h compared to 4 h. Altogether, these results indicate that both sigB expression and σ B activity are stochastic within a subpopulation of C. difficile vegetative cells, likely leading to a "bet-hedging" mechanism.

Discussion

In this study, we deciphered the σ B activation signalling pathway in C. difficile (Fig. 7). The partner switching mechanism responsible for σ B activation that involves RsbV and RsbW is functional in C. difficile. This post-translational mechanism described in other firmicutes relies on the phosphorylation state of RsbV that is dependent on the activity of PP2C phosphatases. We identified CD2685/RbsZ as the phosphatase of RsbV in C. difficile.

While the PP2C phosphatase C-terminal domain is highly conserved, the N-terminal sensory domains and the associated signalling pathways vary considerably among species (de Been et al. 2011, Pane-Farre et al. 2017). In B. subtilis and L. monocytogenes, RsbU phosphatases function with the stressosome, a signal integration and transduction hub multi-protein that is stresses not yet identified are indicated by the "flash" symbol. Locations of potential stimulus perception are indicated by "explosion" symbols. The CD2684 essential iron-sulphur flavoprotein might be involved in stress detection or be involved in cellular process that results in RsbZ phosphatase activation. This activation leads to RsbV dephosphorylation that enables its interaction with the RsbW anti-sigma factor. This protein-protein interaction results in σ B release, its association with RNA-polymerase core enzyme and the transcription initiation of its target genes. domain, respectively (Vijay et al. 2000, de Been et al. 2010). Interestingly, a different GAF domain is associated with the phosphatase domain of CD2685 in C. difficile explaining why we choose a specific name for this phosphatase, RsbZ. In B. cereus, the RsbK sensor protein that activates RsbY also contains a GAF domain, which is proposed to sense an intracellular unidentified stress (de Been et al. 2010). In addition to RsbZ, we showed that CD2684, encoded by the gene in operon with rsbZ, also participates in the control of σ B activity through an uncharacterised mechanism (Fig. 7). CD2684 is an essential iron-sulphur protein sharing similarities with FMN-dependent oxidoreductases. Since most of the genes positively controlled by σ B encode proteins involved in oxidative or nitrosative protection (Kint et al. 2017) and because iron-sulphur clusters are among the first targets of oxidizing compounds (Py et al. 2011), we assumed that CD2684 might be a sensor triggering σ B activation when oxidized. However, we were unable to show any induction of σ B activity after O 2 , H 2 O 2 or DEA/NO treatment (N. Kint, unpublished results). In B. subtilis, the gene encoding the PP2C phosphatase RsbP belongs to an operon with rsbQ, encoding an α/β hydrolase. RsbQ interacts with the PAS domain of RsbP and is required for a proper σ B activity (Brody et al. 2001). In addition, it has been proposed that an unidentified metabolite produced by RsbQ leads to σ B activation through the RsbP PAS domain (Nadezhdin et al. 2011). Contrary to RsbP and RsbQ, no interaction was detected between CD2684 and RsbZ using the BACTH system (Table 2). This result suggests that the effect of CD2684 on σ B activity does not involve a direct interaction with RsbZ but rather acts through either another protein or the synthesis of a metabolite. The essentiality of CD2684 in both the wild-type strain and sigB mutant support the idea that this oxidoreductase might be involved in the production of a crucial metabolite for C. difficile. As proposed for the PAS domain of RsbP in B. subtilis (Nadezhdin et al. 2011), this compound could be detected by the GAF domain located in the N-terminal part of RsbZ modulating its phosphatase activity (Fig. 7). Unfortunately, we did not achieve the production of a stable RsbZ protein lacking the GAF domain preventing us to study the role of this domain in C. difficile. Further studies are needed to characterize the function of the essential CD2684 and to elucidate its exact role in σ B signalling pathway.

While σ B and its regulatory proteins (RsbV and RsbW) are absent in most of the species of the Clostridiaceae family, almost all the species belonging to the Clostridia Cluster XI now reclassified in the Peptostreptococcaceae family harbour the 5-gene sigB operon organisation found in C. difficile (Fig. S5). Strikingly, these species also harbour a genetic Growth curves of the 630∆erm + P tet -rsbV, 630∆erm + P tet -rsbW, 630∆erm ∆rsbZ + P tet -rsbZ, 630∆erm + P tet -rsbZ and 630∆erm ∆rsbZ + P tet -rsbZ∆pho in presence of different concentration of aTc. These data are the mean of 4 independent experiments. locus encoding both a PP2C phosphatase associated to a GAF domain and an FMN oxidoreductase sharing high level of identity with CD2684 (Fig. S5). Conversely, another class of PP2C phosphatase, which does not contain a GAF domain, is present in species belonging to Hungateiclostridiaceae, Eubacteriaceae and Tissierellaceae families. In addition, the corresponding gene is not genetically associated with a gene encoding a CD2684 homolog (Fig. S5). These differences strongly suggest the existence of two distinct σ B activation pathways in Clostridiales, one specific to the Peptrostreptococcaceae and one specific to other families (Fig. S5). However, the genomic organization as well as the smaller set of proteins involved in the control of σ B activity could reflect a more ancestral general stress response activation process in Clostridiales than in Bacillales as proposed for the sporulation process (Paredes et al. 2005, Galperin et al. 2012).

A major breakthrough of our study is also the absence of an auto-regulation of sigB expression in C. difficile. Indeed, in other firmicutes, σ B activation leads to an increased sigB transcription through an internal σ B -dependent promoter located upstream of rsbV, rsbW and sigB resulting in a positive feedback loop (Kalman et al. 1990, Becker et al. 1998, Senn et al. 2005). Since sigB is transcribed only from a σ A -dependent promoter in C. difficile, there is no positive feedback loop increasing sigB expression or σ B activity. In C. difficile, strains overexpressing sigB or genes such as rsbV and rsbZ showed a growth defect likely due to an over-activation of σ B . Interestingly, B. subtilis growth is also affected when sigB expression or σ B activity is higher than the physiological level, since ∆rsbW and ∆rsbX mutants form pinpoint colonies (Kalman et al. 1990, Benson and Haldenwang 1992, Boylan et al. 1992). In B. subtilis and L. monocytogenes, rsbX, which clusters with sigB, encodes a PP2C phosphatase responsible for resetting the stressosome involved in the detection of environmental stresses activating σ B (Marles- Wright et al. 2008, Marles-Wright andLewis 2008). This reset restricts the positive feedback loop and enables σ B signalling pathway to return to its initial state, avoiding toxicity due to an excess of active σ B (Voelker et al. 1997, Xia et al. 2016). The absence of a positive feedback loop on the transcription of sigB operon in C. difficile limiting its expression might explain the absence of any RsbX homolog in this bacterium.

Most of the RsbV protein detected in our Phos-Tag experiment was dephosphorylated in wild-type strain after 4 h of growth (Fig. 4D). Even if we cannot rule out the inability of RsbW to phosphorylate all produced RsbV proteins, this result supports the existence of a σ B activity during the exponential phase. It is worth noting that in S. aureus, a significant proportion of RsbV is dephosphorylated during the exponential growth phase, leading to σ B (1) 630∆erm ∆rsbZ + P tet -rsbZ, (2) 630∆erm ∆rsbZ + P tet -rsbZ-HA and (3) 630∆erm ∆rsbZ + P tet -rsbZ∆pho-HA. Western blot using an antibody raised against the HA tag. Loading control was realized using a 12 % acrylamide gel followed by Coomassie staining.

Figure S3

: Genetic organisation of rsbZ locus. Organisation of rsbZ locus. Gene function is indicated below each gene. TSS mapping experiment indicated the presence of a σ F -dependent promoter upstream of CD2686 (Soutourina et al, 2013, Saujet et al, 2013). 5' Race experiments were realised with NK134 annealing in CD2684 and sequenced with IMV830 annealing in rsbZ to obtain the TSS (see Experimental procedures). The -35 and -10 boxes as well as the TSS are indicated in bold.

Annexes 215 activation and to the transcription of some σ B -targets (Giachino et al. 2001, Senn et al. 2005, Pane-Farre et al. 2009). We provided evidences that σ B is already active during the exponential phase without external stress exposure in C. difficile through the monitoring of P CD1474 -SNAP Cd reporter fusion by fluorescence microscopy. However, we observed a difference in the percentage of cells expressing sigB and CD1474 that could result from a weak sigB expression level in C. difficile that might be under the fluorescence detection threshold. This idea is supported by the high level of sigB overexpression observed in the strain harbouring P tet -sigB in the presence of only 25 ng.ml -1 of aTc (400-fold, Fig. 2C) while sigB was only overexpressed 10-fold when its expression was under the control of its own promoter (Table 1). Heterogeneous σ B activity has also been observed in L. monocytogenes in stressed and non-stressed cells (Utratna et al. 2012, Utratna et al. 2014, Guldimann et al. 2017). This heterogeneity in σ B activity might lead to a specialised sub-population resulting in "bet-hedging" strategy that would maximize the survival chances in adverse environments (Eldar andElowitz 2010, Guldimann et al. 2017). Interestingly, the existence of specialised sub-population has been recently reported for toxin expression in C. difficile. During stationary phase, tcdA expression is indeed governed by σ TcdR through a bistable switch and is also observed in sporulating cells indicating that toxin production and sporulation are not mutually exclusive programs (Ransom et al. 2018a). Conversely, sub-populations triggering stress response and sporulation might be different since σ B represses sporulation while it does not control toxin production (Kint et al, 2017). We can assume that following spore germination, a rapid σ B activation in a sub-population of vegetative cells during exponential growth phase might prepare C. difficile to face the stressful conditions encountered in the gut.

Since the colonization fitness of the sigB mutant is highly impaired in dixenic mice (Kint et al. 2017), it is tempting to speculate that this preadapted sub-population might be important for an efficient gut colonisation by C. difficile. Serial dilutions of the different strains were spotted on TY plates. Plates were incubated 64 h in presence of 0.4 % of O 2 for low O 2 tolerance assay (A) or 24 h at 37°C for NO tests (B). Growth controls were performed with plates incubated in anaerobiosis atmosphere or supplemented with NaOH for low O 2 tolerance assays and NO tests, respectively. C) Effect of CD0007, CD0008 or CD0007-08 deletion on the expression of sigB operon genes and σ B target genes at the onset of the stationary phase. Fold change of the differential expression of genes belonging to sigB operon, to rsbZ operon and of σ B target genes involved in oxidative and nitrosative stress management in 630∆erm ∆CD0007 (black), 630∆erm ∆CD0008 (light gray) and 630∆erm ∆CD0007-08 (dark gray) compared to 630∆erm. Pictures are representative results of 3 independent experiments and qRT-PCR results are the mean with the standard deviation of 3 independent experiments.

were then directly introduced into the pMTLSC7315 plasmid using the Gibson Assembly® Master Mix (Biolabs). The sequences of the resulting plasmids were verified by sequencing.

Plasmids introduced in HB101/RP4 E. coli strain were transferred by conjugation into C. difficile 630∆erm strain. Transconjugants were selected on BHI plates supplemented with Tm and C. difficile selective supplement (SR0096, Oxoid). Isolation of faster growing singlecrossover integrants was performed by serially restreaking on BHI plates supplemented with Cfx and Tm and confirmed by PCR. Single-crossover integrants were then restreaked on C. difficile minimal medium (CDMM) plates supplemented with fluorocytosine (50 µg.ml -1 ) allowing the isolation of double crossover events (Cartman et al. 2012). For CD2684 conditional mutant, aTc (100 ng.ml -1 ) was added to the CDMM plates to maintain CD2684 expression. To confirm the loss of the plasmid, fluorocytosine resistant clones were restreaked on BHI plates supplemented with Cfx and Tm supplemented with aTc for CD2684 conditional mutant. The Tm sensitive clones were checked by PCR for the presence of the expected deletion or for the insertion of P tet element for CD2684 conditional mutant.

For construction of plasmids overexpressing the different genes, fragments were PCR amplified using DNA from the 630∆erm strain. For each construction, a PCR fragment containing the ribosome binding site as well as the gene was cloned into pDIA6103 between the StuI and BamHI sites. For the construction of the strain expressing rsbZ∆pho, we PCR amplified rbsZ gene lacking the part corresponding to the amino acids 336 to 587 containing the PP2C phosphatase domain. In the resulting plasmids, the genes (sigB, rsbW, rsbV, rsbZ or rsbZ∆pho) were expressed under the control of P tet promoter. To construct the translational fusions coding for σ B -, RsbV-and RsbZ∆pho-HA-tagged proteins, plasmids pDIA6325,

Cell lysis, Phos-Tag™ and Western-immunoblot

Whole cell lysates were prepared as previously described (Fagan and Fairweather 2011).

Briefly, cultures of C. difficile were harvested by centrifugation at 5000 r.p.m for 10 min at 4°C and the pellets were frozen at -20°C. Bacteria were thawed and resuspended to an Annexes 218 OD 600nm of 40 in PBS containing 0.12 µg.ml -1 of DNase I. For analysis by SDS-PAGE, an equal volume of 2X SDS sample buffer was added to protein samples (Laemmli 1970). SDS-PAGE and Western immunoblotting were carried out using standard method using anti-HA antibody. Phos-tag™ experiments followed by western-immunoblot were performed according to the manufacturer's instruction. Briefly, after cell lysis, cell extracts were run on a 12 % SDS-PAGE supplemented with 50 µM of Phos-Tag™ acrylamide (AAL-107M, Wako) and 100 µM of MnCl 2 . After gel running, the gel was soaked with agitation three times in fresh transfer buffer supplemented with 10 mM of EDTA for 10 min. Gel was then soaked 10 min in fresh transfer buffer without EDTA before gel transfer to the PVDF membrane.

Western immunoblotting using anti-HA antibody was then carried out using standard method.

Spotted dilution assays

O 2 tolerance assays and NO stress assays were performed as previously described (Kint et al. 2017). Five microliters of serial dilutions (from 10 0 to 10 -5 ) of C. difficile strains grown for 8 h in the TY medium were spotted on TY plates supplemented i) with 25 ng.ml -1 of aTc and 0.05% of taurocholate and incubated in the presence of 0.4 % O 2 tension; or ii) with 25 ng.ml - 1 aTc containing 250 or 500 µM of DEA/NO. The last dilution allowing growth was recorded after incubation at 37°C for 64 h for the O 2 tests and for 24 h for the NO stress tests.

Transcriptional SNAP Cd fusions

To construct SNAP fusions, we used pFT47, a pMTL84121 plasmid derivative containing a synthetic SNAP Cd sequence (Pereira et al. 2013, Cassona et al. 2016). To construct CD1474-SNAP Cd transcriptional fusion, a 200 bp PCR product containing the promoter region of this gene (position -206 to +19 from the TSS) was amplified using genomic DNA and primer pairs CF19/CF20 and inserted between the EcoRI and XhoI sites of pFT47 to obtain PDIA6457. The insert sequence was checked by sequencing. The plasmid was transferred by conjugation into C. difficile 630∆erm strain and sigB, ∆rsbV or ∆rsbZ mutant giving CDIP661, CDIP668, CDIP728 and CDIP953 strains, respectively (Table S1). To construct sigB-SNAP Cd transcriptional fusion, plasmid pDIA6325 used for complementation containing both the sequence located upstream of CD0007 (position -120 to +50 from the TSS) and sigB gene was digested by XhoI and StuI to remove the sigB gene and keep the σ A dependent promoter located upstream CD0007. SNAP Cd gene was PCR amplified using pFT47 DNA and primers pairs NK155/NK156. The DNA fragment was digested by DpnI to remove pFT47 plasmid trace and inserted between the XhoI and StuI sites of pDIA6325 to obtain Annexes 219 pDIA6589. The resulting plasmid was then transferred into 630∆erm strain and sigB mutant, giving CDIP843 and CDIP845 strains, respectively.

Fluorescence microscopy and image analysis

Overnight culture of CDIP843, CDIP845, CDIP661 and CDIP668 strains were diluted at an OD 600nm of 0.05 into 5 ml of BHI. For SNAP labelling, 200 µl of culture were mixed with 250 nM SNAP-Cell TMR-Star (New England Biolabs) and incubated 30 min at 37°C in the dark at each time point (4, 10, 24 and 48 h) (Pereira et al. 2013, Cassona et al. 2016). After labelling, cells were collected by centrifugation (4000 g for 3 min) and washed 3 times with 1 ml of PBS. For fluorescence microscopy, cells were mounted on 1.7 % agarose coated glass slides to keep the immobilized cells, and to obtain sharper images (Glaser et al. 1997). The images were taken with exposure times of 200 ms for autofluorescence and 500 ms for SNAP.

Cells were observed on a Nikon Eclipse TI-E microscope 60x Objective and captured with a CoolSNAP HQ2 Camera. For quantification of the SNAP-TMR Star signal resulting from transcriptional fusions, the pixel intensity was measured, and corrected by subtracting the average pixel intensity of the background. Images were analysed using ImajeJ.

Bacterial adenylate cyclase two hybrids system (BACTH) and ß-galactosidase assays

Protein-protein interactions were analysed by the BACTH system (Euromedex). All the procedures for plasmids constructions and E. coli co-transformations were performed according to the manufacturer's instruction. Briefly, this system exploits the catalytic domain of the adenylate cyclase from B. pertussis (CyA) that is composed of two different fragments T25 and T18. When T25 and T18 are separated, CyA is not active. However, when these fragments are fused with interacting proteins, the heterodimerisation of these hybrid proteins allows a functional complementation and so production of cAMP leading to lacZ gene expression. The open reading frames of rsbV, rsbW, sigB or rsbZ were cloned into plasmids allowing fusions with T18 or T25 either in C-terminal or N-terminal position. After verification by sequencing, the different association of T18 and T25 fusion plasmids were cotransformed in E. coli strain BTH101 (Table S1). Co-transformation with the T25 and T18 expressing plasmids served as a control for the basal β-galactosidase activity. The βgalactosidase activity was measured after an overnight culture of co-transformants in LB medium in the presence of 0.5 mM of isopropyl-β-D-thiogalacto-pyranoside. 100 µl of overnight culture was mixed with 400 µl of M63 medium and 500 µl of Z buffer (Na 2 HPO 4 Annexes 220 0.06 M, NaH 2 PO 4 0.04 M, KCl 0.01M, MgSO 4 0.001 M, DTT 0.001 M and SDS 0.0037 %).

After addition of 20 µl of chloroform, the tubes were mixed exactly 10 s. Tubes were preincubated 5 min at 28°C and 200 µl of pre-heated ONPG (4 mg.ml -1 ) was added. When the yellow coloration appeared, 500 µl of Na 2 CO 3 (1 M) was added to the tube to stop the reaction. After elimination of cell fragments, the OD 420nm was measured.

RNA extraction, quantitative RT-PCR analysis and 5' RACE experiment

To test the expression of σ B target genes, overnight cultures of the different strains were diluted to an OD 600nm of 0.05 in 10 ml of TY medium supplemented with 25 ng.ml -1 aTc when needed. Cells were harvested after 10 h of growth. 630∆erm strain cells were harvested after 4, 10, 16, 20 or 24 h of growth. To test the effect of sigB overexpression, CDIP219 and CDIP393 (Table S1) grown in 10 ml of TY medium without aTc or with 25 ng.ml -1 or 200 ng.ml -1 of aTc were harvested after 4 h of growth. To study the effect of CD2684 depletion, overnight culture of CD2684 conditional mutant was made in presence of 10 ng.ml -1 of aTc to allow a growth. After dilution to an OD 600nm of 0.05 in 10 ml of TY medium without or with 10 ng.ml -1 of aTc, cells were harvested after 4 or 10 h of growth. RNA extraction, cDNAs synthesis and real-time quantitative PCR were performed as previously described (Saujet et al. 2011, Soutourina et al. 2013). The quantity of cDNAs of a gene was normalised to the quantity of cDNAs of the pgi and ccpA genes. The relative change in gene expression was recorded as the ratio of normalised target concentrations (the threshold cycle [∆∆C T ] method) (Livak and Schmittgen 2001). 5' RACE experiment was realised with the 5' RACE system for Rapid Amplification of cDNA Ends kit (Invitrogen) according to the manufacturer's instructions. Briefly, a first strand cDNA was synthesised from total RNA extracted after 10 h of growth in TY using Superscript II and primer NK134 annealing in CD2684. A C-tail was then added to the 3' end of the cDNA using TdT and dCTP. The resulting fragment was amplified by the Abridged Anchor Primer and NK31 annealing in rsbZ. The resulting fragment was then sequenced by IMV830 annealing in rsbZ to define the TSS. The erm gene encodes the erythromycine resistance cassette. The cat gene encodes the chloramphenicol resistance cassette leading to resistance to chloramphenicol, Cm R and to thiamphenicol, Tm R . Amp R means that the plasmid harbors a resistance cassette responsible for ampicillin resistance.

Table S3. Oligonucleotides used in this study.

a Restriction sites and tales used for Gibson assembly clonning are underlined. Total RNAs were extracted from the wild-type and CD1065 mutant strains after 14 h (for σ Eor σ F -target genes) or 24 h (for σ G -or σ K -target genes) of growth in SM medium. The results presented correspond to the mean of at least 3 independent experiments.

Primer

Abstract

The strict anaerobe and sporogenic Clostridium difficile is the most common cause of antibiotic-associated diarrhoea. Spores have a central role in the C. difficile infectious cycle: resistance outside the host, persistence of bacteria during infection and transmission of the disease. When ingested by the host, in the presence of certain bile salts, C. difficile spores germinate in the colon to form vegetative cells that secrete toxins and cause the symptoms of infection.

Vegetative cells are exposed to several stresses, among them, reactive oxygen/nitrogen species (ROS/RNS) produced by the host immune system during inflammation. Furthermore, although the intestinal tract is regarded as mainly anoxic, low oxygen (O 2 ) tensions are present in the large intestine and tend to increase after antibiotic treatments. To survive to this hostile oxidative environment, C. difficile had to develop mechanisms of protection, detoxification pathways and repair systems. The alternative sigma factor of the general stress response, σ B , plays an important role in the protection of C. difficile against the different stresses the bacterium is facing inside the host including O 2 and ROS. Among the genes positively controlled by σ B , we identified genes encoding proteins likely involved in O 2 tolerance and ROS detoxification, such as the two reverse rubrerythrins (revRbrs), CD1524 and CD1474, and the flavodiiron proteins (FDPs), CD1157 and CD1623. We showed that the two revRbrs and the FDP CD1623 are expressed in a σ B -dependent manner and heterogeneously in a cell population. A dual genetic control was demonstrated for the FDP encoding gene CD1157, through σ A -and σ B -dependent promoters. We demonstrated that CD1157 displays O 2 -reductase activity while the two revRbrs have both H 2 O 2 -and O 2reductase activity, being H 2 O 2 the preferred substrate. Moreover, we showed that a double ∆CD1474-∆CD1524 mutant and a CD1157 mutant are less tolerant to low O 2 tension and that the CD1623 mutant is more sensitive to air exposure. These findings demonstrate a key role for these proteins in O 2 tolerance and oxidative stress response in C. difficile.

During the infection cycle, in the colon some vegetative cells are transformed into spores. The spore surface layers, the coat and exosporium, enable the spores to resist physical and chemical stress. However, little is known about the mechanisms of their assembly. In this work, we characterized a new spore protein, CotL, which is required for the assembly of the spore coat. We showed that the cotL gene is expressed in the mother cell compartment under the dual control of the sporulation sigma factors, σ E and σ K . CotL was localized in the spore coat, and the spores of the cotL mutant had a major morphologic defect at the level of the coat/exosporium layers. Therefore, the mutant spores contained a reduced amount of several coat/exosporium proteins and a defect in their localization in sporulating cells. Finally, cotL mutant spores were more sensitive to lysozyme and were impaired in germination, a phenotype likely associated with the structurally altered coat. Collectively, these results strongly suggest that CotL is a morphogenetic protein essential for the assembly of the spore coat in C. difficile. Keywords: Clostridium difficile, Spore, Coat, Morphogenesis, CotL, Oxygen, ROS, detoxification, Rubrerythrins, Flavodiiron proteins

  Fight or flight?

Figure 1 :

 1 Figure 1: Morphology of Clostridium difficile. A) Phase-contrast microscopy image of C. difficile culture. B) C. difficile colonies on plate.

Figure 3 :

 3 Figure 3: Potential mechanisms involved in colonization resistance against C. difficile in the gut. Direct inhibition of C. difficile by the gut microbiota can occur via the conversion of nutrients or host metabolites to compounds that are inhibitory to C. difficile, production of compounds with bacteriostatic activity (e.g. bacteriocins) and competition for nutrients. Indirect control of C. difficile expansion can occur via the stimulation of the host immune system through the production of microbial-associated molecular patterns (MAMPs). Detection of MAMPs by the host induces the activation of an host immune cascade and the production of innate (e.g. antimicrobial peptides) or adaptive (e.g. IgA) immune effectors. SCFA, short-chain fatty acids. Adapted from Britton and Young, 2012.

Figure 4 :

 4 Figure 4: Effect of bile acid metabolism on the life cycle of C. difficile. The liver synthesizes bile acids from cholesterol and secretes them as conjugated bile acids. Ileum gut microbiota produces bile salt hydrolases that deconjugate bile acids to primary bile acids, chenodeoxycholate and cholate. Primary bile acids serve as substrates for 7-dehydroxylases, made by a small subset of clostridial organisms such as C. scindens, which generate secondary bile acids lithocholate and deoxycholate. Chenodeoxycholate and secondary bile acids are toxic to vegetative cells of C. difficile (red lines), while cholate derivatives promote germination when combined with amino acid co-germinants like glycine (dotted line). Shen et al., 2015.

  a. Mechanism of action of TcdA and TcdB

Figure 6 :

 6 Figure 6: C. difficile pathogenicity locus (PaLoc). Both tcdA and tcdB genes are encoded on the 19.6-kb pathogenicity locus (PaLoc), together with tcdR, tcdC and tcdE genes.TcdR is an alternative sigma factor required for the transcription of tcdA, tcdB and tcdR itself. TcdC is an anti-sigma factor which sequesters TcdR preventing the transcription of the toxin genes. TcdE is a "holin" type bacteriophage protein required for toxin secretion.

  al.

Figure 7 :

 7 Figure 7: Regulation of the expression of PaLoc genes. In the presence of nutrients (amino acids, GTP or PTS sugars), the transcriptional regulators, CodY and CcpA, repress the expression of toxin genes by binding to the promoter regions of the PaLoc genes. Proline, glycine and cysteine also have a negative influence on toxin production, but the molecular mechanisms of this repression are not yet fully understood. σ H and the master regulator of sporulation Spo0A repress indirectly the expression of the toxin genes. The regulator RstA indirectly control the expression of the toxins by repressing the transcription of σ D.

Figure 8 :

 8 Figure 8: Schematic representation of the mechanism of action of CDT. The monomeric form of CDTb binds to its cell surface receptor, LSR. Thereafter, CDTb undergoes proteolytic activation and oligomerization on the cell surface. This process might also occur before binding. The oligomeric prepore binds CDTa, and the receptor-CDT complex is subsequently internalized into cells. As the endosome matures, the acidic pH triggers conformational changes in CDTb, resulting in membrane insertion and formation of a pore through which CDTa is translocated into the cytosol. Translocation of CDTa is facilitated by heat shock protein 70 (Hsp70), Hsp90 and cyclophilin A (CypA). In the cytosol, CDTa ADP-ribosylates actin at arginine-177. ADP-ribosylation of actin results in the eventual breakdown of the actin cytoskeleton and cytopathic cell rounding, and the formation of microtubule-based protrusions that enhance bacterial adherence. Chandrasekaran and Lacy 2017.

Figure 9 :

 9 Figure 9: Organization of the B. subtilis sigB operon, genes encoding different regulators and model of σ B activation. A) Open reading frames (arrows) and transcriptional start sites with promoters are indicated. Structurally and functionally conserved domains are color coded. Abbreviations: STAS (domain found in sulfate transporters and anti-sigma factor antagonists), Sigma (sigma factor), HATPase (kinase domain of two-component transmitter proteins), PP2C (Mn 2+ /Mg 2+ -dependent protein phosphatase 2C), PAS (Per-ARNT-Sim signal sensor domain implicated in the perception of redox, voltage, and/or, light signals). Adapted from Hecker et al., 2007. B) Partner switching mechanism involved in σ B activation.In unstressed cells and during the exponential growth phase, the anti-sigma factor, RsbW, phosporylates the anti-anti sigma, RsbV, and sequesters σ B rendering it inactive. During stationary growth phase of after stress exposure, the non-phosphorylated form of RsbV interacts with RsbW, allowing σ B release, its association with the RNA polymerase and transcription of its targets genes.

  stress response and the role of σ B

Figure 10 :

 10 Figure 10: Model of the signal transduction leading to the activation of σ B in B. subtilis. A) Energy-stress mediated activation of RsbP phosphatase. B) Environmental stress-mediated activation of RsbT and its subsequent release from the stressosome.

  Figure 11: C. difficile σ B has a pleiotropic role. Red and green arrows indicate respectively positive and negative control of σ B under the different processes. σ B positively regulates the expression of several genes involved the management of stresses find by C. difficile during gut colonization: e.g. oxygen tolerance, oxidative and nitrosative stress, thiol homeostasis, low pH adaptation and antimicrobial peptides resistance.

Figure 12 :

 12 Figure 12: Organization of the C. difficile sigB operon. Open reading frames (arrows) and the transcriptional start site with the promoter recognized by σ A are indicated. The function of proteins encoded by each gene is indicated above each gene. Percentage of identity with the B. subtilis proteins is indicated below each gene.

Figure 13 :

 13 Figure 13: Enzymatic systems for detoxification of oxygen (O 2 ) and reactive oxygen species (ROS). SOD (superoxide dismutase), CAT (catalase), SOR (superoxide reductase), Pxd (peroxidase), Rbr (rubrerythrin), revRbr (reverse rubrerythrin), FDP (flavodiiron protein) and O 2 R (respiratory O 2 reductase) are indicated.

Figure 14 :

 14 Figure 14: Schematic representation of a Rubrerythrin (Rbr) and a reverse Rubrerythrin (revRbr) domains. Rbr contains non-sulfur diiron center in the N-terminal part and a rubredoxin-type [FeCys 4 ] center, located at the C-terminal part. In a revRbr, the position of the two domains is switched.

Figure 15 :

 15 Figure 15: The FDP protein family. This scheme assumes a FDP family classification according to the known or proposed domain structure (deduced from their gene translated amino acid sequences). Structural domains are color coded. Adapted from Folgosa et al., 2018.

Figure 17 :

 17 Figure 17: Oxygen and ROS detoxification system in C. acetobutylicum. Scheme of the NADH-dependent O 2 , O 2-and H 2 O 2 detoxification complex composed of NADH-rubredoxin oxidoreductase (NROR), rubredoxin (Rd), flavoprotein (Fpr), desulfoferrodoxin (Dfx) and rubrerythrin (Rbr). Adapted from[START_REF] Kawasaki | O2 and reactive oxygen species detoxification complex, composed of O2-responsive NADH:rubredoxin oxidoreductase-flavoprotein A2-desulfoferrodoxin operon enzymes, rubperoxin, and rubredoxin, in Clostridium acetobutylicum[END_REF]

Figure 18 :

 18 Figure 18: Oxygen tension in the colon before and after antibiotic treatment. A) O 2 tension and metabolic reactions in a healthy colon. The butyrate produced by the microbiota, and in particular by certain clostridia, is oxidized to CO 2 by mature colonocytes to produce energy (ATP). The consume of O 2 by mature colonocytes renders the epithelial surface hypoxic. B) Metabolic change and increase in O 2 pressure after antibiotic treatment. After antibiotic treatment, the intestinal microbiota is affected resulting in a decrease in butyrate production. In this case, mature colonocytes ferment glucose into lactate to obtain ATP. The use of this metabolism increases oxygenation of the epithelial surface. The resulting increased diffusion of O 2 into the intestinal lumen drives an aerobic expansion of organisms such as of Enterobacteriaceae. The color scale shown at the left bottom of the panel indicates the O 2 concentration signified by the coloring scheme of the schematic.

Figure 19 :

 19 Figure 19: Formation of spores by endospore-forming bacteria and compartimentalization of gene expression. The main morphogenetic stages of the sporulation process are represented in the scheme. The compartment and main periods of activity of the sporulation σ F , σ E , σ G and σ K sigma factors are indicated in the respective morphological stages. Adapted from[START_REF] Errington | Regulation of endospore formation in Bacillus subtilis[END_REF]

Figure 20 :

 20 Figure20: Activation of Spo0A in B. subtilis. Spo0A phosphorylation is controlled by a phosphorelay that depends on the activity of several kinases and other intermediaries. Interactions that ultimately lead to Spo0A phosphorylation are indicated in black, while the ones working against it are in red. The KinA and KinB kinases respond to yet not known signals by autophosphorylating and subsequently transferring phosphate to Spo0F. Spo0F phosphorylates Spo0B, which passes phosphate on to Spo0A. A third kinase, KinC, bypasses the phosphorelay and acts directly on Spo0A. Phosphatases, including RapA, B and E and Spo0E, and kinase inhibitors (Sda) act at various steps in the pathway to directly or indirectly antagonize Spo0A phosphorylation.

Figure 21 :

 21 Figure 21: The sporulation network of B. subtilis. Temporal progression of sporulation is shown from top to bottom. Boxes represent the predivisional cell (PDC), mother cell (MC) and forespore (FS), the two cell compartments formed after asymmetric division. Membranes separating the FS and the MC are indicated (OFM: outer forespore membrane; IFM: inner forespore membrane). Sigma factors are shown colored and bold, with the precursor proteins indicated as pro-σ E /σ K , transcriptional regulators are shown in purple and anti-sigma factors in brown. Proteins associated with the membranes or located into the intermembrane space are in square. Black solid arrows indicate activation at the transcriptional level. Red arrows indicate the intercellular signal pathways involved in the sigma factor activation.

Figure 25 :

 25 Figure 25: B. subtilis spore germination. Germinants such as L-alanine (or L-valine) interact with the GerA germinant receptor complex. The SpoVA channel is then activated and it releases Ca-DPA from the spore core. Released of Ca-DPA activates the CwlJ cortex hydrolase triggering cortex hydrolysis. OM, GCW and IM correspond to outer membrane, germ cell wall and inner membrane, respectively.

Figure 27 :

 27 Figure 27: Structure of a bacterial spore. A) The main layers of the spore are represented. The most external layer, either crust or exosporium, is not present in all spore formers. Transmission elecron microscopy of a B. subtilis spore (B) and a C. difficile spore (C). Co, Cx, IC, OC, Cr and Ex are abbreviations for core, cortex, inner coat, outer coat, crust and exosporium, respectively (B. subtilis picture was adapted from McKenney et al., 2010).

Figure 28 :

 28 Figure 28: The B. subtilis coat interaction network. Morphogenetic proteins are indicated on the upper part of the figure and are located where their main function is exerted. The hexagon represent the LysM domain. Arrows indicate the interactions between the different morphogenetic coat and crust proteins Examples of coat proteins, which proper location is dependent on the respective morphogenetic protein are indicated on the bottom of the figure. Assembly of each layer may be driven by the multimerization of the indicated morphogenetic protein. Polymerization of a morphogenetic protein would, presumably, create the necessary binding sites for each of the individual coat proteins that make up each layer. It remains to be determined whether and how the proteins of the individual layers interact with adjacent layers and how adjacent layers interact to form the mature spore coat. Adapted from Henriques and Moran 2007 and McKenney et al., 2013.

Figure 29 :

 29 Figure 29: Model of spore coat assembly during B. subtilis sporulation. In the left column, are listed the stages of sporulation. The center column contains diagrams of spore coat morphogenesis. Layers of the spore coat are color coded (cyan = basement layer; yellow = inner coat; blue = outer coat; maroon = crust). In the right column, are listed the stages of spore coat assembly. DPA, dipicolinic acid. Driks 2016.

  Figure 30: Thin-sectioned spores of B. anthracis (A) and B. thurigiensis (B). The main layers of the spore are represented. Co, Cx, IC, OC, C and Ex are abbreviations for core, cortex, inner coat, outer coat, coat and exosporium, respectively. The exosporium is the outermost protective layer of the spore. Exosporium contains hair-like projections mainly composed by the BclA protein. Exosporium is separated from the rest of the spore by a interspace. B. anthracis picture was adapted from Stewart 2015; B. thurigiensis picture was adapted from Gerhardt et al., 1976.

  . It is important to note that C. difficile CotE bears no sequence or functional homology to the B. subtilis outer coat morphogenetic protein CotE (McKenney et al. 2013). Discrimination of spore proteins localization within the coat or the exosporium of C. difficile spores was difficult until the establishment of methods that allow to separate these two layers. The development of exosporium removal methods, that mantain the spore coat intact, combined with mass spectometry allowed the study of the proteomic composition of the exosporium of C. difficile spores (Escobar-Cortes et al. 2013, Diaz-Gonzalez et al. 2015).

  C. difficile, three paralogs of the B. anthracis collagen-like glycoprotein (i.e., BclA) are encoded in the genome of the 630 strain (i.e., bclA1 [CD0332], bclA2 [CD3230], and bclA3 [CD3349]). These genes are expressed in the MC under σ K control (Pizarro-Guajardo et al. 2014). The BclA proteins contain three domains: an N-terminal domain that localizes BclA to the exosporium, a collagen-like domain, and a C-terminal domain facing to the surface of the exosporium (Pizarro-Guajardo et al. 2014). BclA1 undergoes post-translational cleavage and forms high molecular complexes with other spore proteins, suggesting that it is assembled in the spore surface through cleavage and/or crosslinking to other exosporium proteins (Pizarro-Guajardo et al. 2014). No homologues of ExsFA and ExsFB proteins involved in the localization of BclA and the stability of the exosporium (Sylvestre et al. 2005) are encoded in the C. difficile genome. Additionally no conserved motifs were found in the N-terminal domain of all three BclAs suggesting that the assembly mechanism is different in C. difficile

Figure 31 :

 31 Figure 31: Schematic representation of proteins of the spore surface layers of C. difficile. Morphogenetic proteins are indicated on the upper part of the figure and are located where their main function is exerted. The hexagon represent the LysM domain. CotA and CotB are proposed to be located in the inner spore coat layers, whereas BclA and Cde proteins are in the exosporium. Adapted from Calderon-Romero et al., 2018.

(

  Lawley et al. 2009b, Permpoonpattana et al. 2011b, Abhyankar et al. 2013, Diaz-Gonzalez et al. 2015). In fact, only 25% of the C. difficile coat proteins have homologs in the spore former model B. subtilis(Henriques and Moran 2007, Permpoonpattana et al. 2011b, Permpoonpattana et al. 2013). In B. subtilis, the spore surface is primarily composed of proteins produced in the MC under the control of first σ E and later σ K(Henriques and Moran 2007, McKenney et al. 2013). In this bacterium, the formation of the coat layers is governed by the morphogenetic factors SpoVM, SpoIVA, SpoVID, SafA, and CotE(Henriques and Moran 2007, McKenney et al. 2013). In the basement layer of the coat, the SpoVM, SpoIVA and SpoVID proteins are required to start the coat localization around the FS surface.Recruitment of SafA, the inner coat morphogenetic protein, and of CotE, the morphogenetic protein of the outer coat, is dependent on SpoIVA and SpoVID(McKenney et al. 2013). Both SpoVM and SpoIVA homologs are present in C. difficile(Galperin et al. 2012). In this bacterium, SpoIVA was shown to be essential for coat localization around the FS(Putnam et al. 2013). However, contrary to the observed in B. subtilis, SpoVM was found to be dispensable for spore formation/maturation. No homologs of B. subtilis SpoVID, SafA and CotE were found in C. difficile. However, a functional homolog of SpoVID, SipL, was identified in C. difficile. As SpoVID, SipL interacts with SpoIVA and might form a complex Results 75 that drive coat assembly around the FS(Putnam et al. 2013). The morphogenetic proteins for the inner and outer coat layers of C. difficile are still unknown.Among the genes whose expression is controlled by the specific σ factors of MC, it was identified CD1065 (later renamed cotL) encoding CotL protein of unknown function with no homologs in bacilli. In this work, we then demonstrated that CotL is an important morphogenetic factor of the coat layer of C. difficile spores. The cotL gene was expressed in the MC compartment under the dual control of the sporulation sigma factors, σ E and σ K . CotL is present in the spore coat and the cotL mutant spores lack most of the spore coat layers and are more sensitive to lysozyme. Using translational SNAP fusions with several coat and exosporium proteins, we showed that CotL is required for the assembly of these proteins around the FS surface during sporulation. Finally, cotL mutant spores were impaired in germination, a phenotype likely to be associated with the structurally altered coat. Our results bring insights on a new morphogenetic partner important for coat assembly of C. difficile spores.

Figure 1 :

 1 Figure 1: Mother cell specific expression of CD1065. A) Promoter region of the CD1065 gene. The mapped transcriptional start sites (+1, yellow), and the -10 and -35 promoter elements corresponding to the consensus for the promoters recognized by σ E (P1 blue boxes) and σ K (P2 red boxes) are indicated. The possible ribosome-binding site (RBS) and the start codon of CD1065 are indicated as grey and green boxes, respectively. B, C and D) Temporal and cell type-specific expression of CD1065 during sporulation. Microscopy analysis of the cells carrying P σE/σK(CD1065) -SNAP Cd (B), P σE(CD1065) -SNAP Cd (C) or P σK(CD1065) -SNAP Cd (D) transcriptional fusions in the wild-type 630Δerm strain and in the sigE or sigK mutant. The cells were collected after 24 h of growth in liquid SM, stained with TMR-Star and the membrane dye MTG, and examined using fluorescence microscopy. The merged images showed the overlap between the TMR-Star (red) and the MTG (green) channels. The panels are representative of the expression patterns observed for different stages of sporulation and are ordered from early to late stages for the wild-type strains. For the sigE and sigK mutants, a morphological stage characteristic of each mutant is indicated. The data shown are representative of at least three independent experiments. The numbers represent the percentage of cells at the indicated stage expressing the reporter fusion. For each stage, the number of cells analysed (n) is as follows: asymmetric division, n=20-30; engulfment, n=40-60; cortex/coat maturation, n= 90-100. For sigE and sigK mutants, n= 30-40. Scale bar, 1 µm.

Figure 2 :

 2 Figure 2: The CD1065 mutant is able to sporulate. Phase-contrast microscopy of sporulating cells (A) and purified spores (B) of the wild-type, CD1065 mutant and complemented strains after 24 h of growth in liquid SM. Yellow arrows mark the mature phase-bright spores; red arrows indicate the phase-grey spores, and green arrows indicate the immature, phase-dark forespores. In panel A, the numbers represent the percentage of sporulating cells. Scale bar, 2 µm. C) Spore length and width of the wild-type and CD1065 mutant strains. The red line indicates the median. n=300. Statistical analysis was conducted using a Wilcoxon test, *P <0.05. D) Counts (CFUs/mL) of total cells (black) and heat-resistant spores (grey) for the wild-type, CD1065 mutant and complemented strains grown for 24 h in SM and plated on BHI with 0.1% TA. Data represent the average of four biological replicates. Statistical analysis was conducted using a Wilcoxon test, *P <0.05.

Figure 3 :

 3 Figure 3: CD1065 mutant spores are defective in the earliest stages of germination. A)Change in the OD 600nm of the spores of the wild-type, CD1065 mutant and complemented strains after the addition of 1% TA. OD 600nm measurements were taken every 5 min. B) CFUs/mL obtained after plating the purified spores on BHI with 0.1% TA. Data represent the average of four biological replicates. Statistical analysis was conducted using a Wilcoxon test, *P <0.05. C) Purified spores incubated in BHI in the presence of 1% TA were fixed in formaldehyde and visualized using phasecontrast microscopy. Scale bar, 2 µm. D) Processing of the SleC in response to TA. The spores were suspended for 20 min in BHI containing 1% TA, and the SleC cleavage was tested by Western blot analysis using anti-SleC antibody. The volume of protein extract applied on the gel was increased three-fold for the second Western blot. Coomassie gel was used as the loading control.

Figure 4 :

 4 Figure 4: CD1065 mutant spores have a defective spore coat layer. A) Thin sections of the wild-type, CD1065 mutant and complemented spores were analysed using TEM as described in the Materials and Methods. Ex, exosporium, Oc, outer coat, Ic, inner coat, Ct, cortex, Cr, core. Scale bars are shown in each figure. The numbers on the bottom of each figure represent the spore dimensions for each strain (length and width) based on the analysis of at least 10 individual spores. Statistical significance was determined using a Wilcoxon test (*P <0.05). B) The thickness of the exosporium, the outer/inner coat layers, and the cortex layer of the spores from the three strains were measured. Error bars denote standard errors of the means of at least 10 individual spores. Statistical significance was determined using a Wilcoxon test (*P <0.05). C) Size of the spore core of the wild-type, the CD1065 mutant and the complemented strains. The length and width of the core was measured. Error bars denote the standard errors of the means of at least 10 individual spores.

Figure 5 :

 5 Figure 5: Lysozyme triggers germination of CD1065 mutant spores. A) Phase-contrast microscopy of the wild-type, CD1065 mutant and complemented spores treated with or without 250 µg/mL of lysozyme for 1 h at 37°C. Scale bar, 2 µm. B and C) CFU/mL obtained of the wildtype, CD1065 mutant and complemented spores untreated (black) and lysozyme-treated (grey) after plating onto BHI agar (B) or BHI agar containing 0.1% TA (C). Data represent the average of the results of at least three independent experiments, and the error bars represent standard errors of the means. Statistical analysis was conducted using a Wilcoxon test, *P <0.05.

Figure 6 :

 6 Figure 6: CD1065 protein is localized in the coat/exosporium layer of C. difficile spores.Western blot analyses of coat/exosporium (3) and decoated spore lysate (1 and 2) fractions from the wild-type, CD1065 mutant and complemented strains. The decoated lysate contains the spore core and cortex layers untreated (1) or lysozyme-treated to separate the cortex layer of the core (2). CotB was used as a control for a coat protein.

Figure 7 :

 7 Figure 7: Subcellular localization of the CotL protein fused to SNAP Cd . Microscopy analysis of C. difficile cells harboring P σE/σK(cotL) -cotL-SNAP Cd (A) or P σK(cotL) --cotL-SNAP Cd (B) translational fusions in the wild-type strain and the sigE or sigK mutant. The cells were collected after 24 h of growth in liquid SM, stained with TMR-Star and examined by phase-contrast (PC) and fluorescence microscopy to monitor the SNAP production. The merged images show the overlap between the TMR-Star (red) and the autofluorescence (green) channels. Wild-type carrying pFT58 was used as the negative control. A schematic representation of the location pattern of the CotL-SNAP Cd protein is shown on the right side of each panel, ordered from early to late stages. Among the cells with signal, the numbers represent the percentage of sporulating cells/spores showing the corresponding pattern of coat protein location. The data shown are representative of three independent experiments. At least 100 cells were analysed for each strain. Scale bar, 1 µm.

Figure 8 :

 8 Figure 8: Fluorescence localization of coat proteins fused to SNAP Cd . Microscopy analysis of the wild-type and cotL mutant cells carrying (A) SpoIVA-and (B) CdeC-SNAP Cd translational fusions. The cells were collected after 24 h of growth in liquid SM, stained with TMR-Star and examined by phase-contrast (PC) and fluorescence microscopy to monitor the SNAP production. The merged images show the overlap between the TMR-Star (red) and the autofluorescence (green) channels. The yellow arrows point the fluorescent signal. A schematic representation of the location pattern of the different proteins fused to SNAP Cd is shown on the right side of each panel. Among the cells with signal, the numbers represent the percentage of sporulating cells/ spores showing the corresponding pattern of coat protein location. The data shown are representative of three independent experiments. For each strain, 100 cells were analysed. Scale bar, 1 µm.

Figure S1 :

 S1 Figure S1: Detection of SNAP production from CD1065-SNAP Cd transcriptional fusion using native or truncated promoters and different backgrounds. The accumulation of SNAP Cd in extracts produced from sporulating cells of the wild-type 630Δerm strain, sigE or sigK mutants harboring P σE/σK(CD1065) -SNAP Cd fusion (A) or the wild-type strain harboring P σE/ σK(CD1065) -SNAP Cd , P σE(CD1065) -SNAP Cd or P σK(CD1065) -SNAP Cd fusions (B). 630Δerm carrying pFT47 was used as a negative control. Proteins in whole cells lysates were resolved by SDS-PAGE and subjected to immunoblot analysis using an anti-SNAP antibody. A section of the corresponding Coomassie-stained gel is shown as a loading control. The position of molecular mass markers (kDa) is shown on the left side of the panels. The experiment was repeated at least three times with different samples.

Figure S2 :

 S2 Figure S2: Construction of the CD1065 mutant in C. difficile. A) Schematic representation of the inactivation strategy for the CD1065 gene using the ClosTron technology. Primers were designed to retarget the group II intron of pMTL0007-CE5 to insert it into the CD1065 gene in a sense orientation immediately after the 123 rd nucleotide of its coding sequence. B) Verification of the L1.LtrB intron integration into the CD1065 gene in the 630Δerm strain. PCRs were performed with DNA of the 630Δerm strain and the CD1065 mutant using primers (CF58/CF59) flanking the insertion site in CD1065 (lane 3), one primer flanking the insertion site in CD1065 (CF58) and the intron primer EBSu (lane 2) or primers (Erm-F/Erm-R) amplifying the erythromycin cassette (lane 1).

Figure S3 :

 S3 Figure S3: Dipicolinic acid (DPA) release by spores of the wild-type, the CD1065 mutant and the complemented strains. Spores of the indicated strains were suspended in PBS supplemented with TA (1%) and glycine (10 mM). After 1 h exposure, the amount of DPA in the germination solution was determined as indicated in the Materials and Methods. The data represent the average and standard deviation from three biological replicates. Statistical significant difference relative to the wild-type was determined using a Wilcoxon test, *P <0.05.

Figure S4 :

 S4 Figure S4: CD1065 inactivation affects the abundance of the major proteins of the spore surface. A) Coat/exosporium extracts from purified spores of the wild-type, the CD1065 mutant, and the complemented strains were loaded on acrylamide gel and stained with Coomassie brilliant blue. Red arrows highlight the major differences in the protein bands. B and C) Western blot analyses using coat/exosporium extracts of purified spores (B) or extracts of sporulating cells (C) using antibodies raised against different spore coat or exosporium proteins (SipL, CotB, CotE, CspC, CspBA, SleC, CdeC and CdeM). Experiments were repeated at least three times with independent samples.

Figure S5 :

 S5 Figure S5: Fluorescence localization of CotB-SNAP Cd . Microscopic analysis of the wild-type and cotL mutant cells carrying a CotB-SNAP Cd translational fusion. The cells were collected after 24 h of growth in liquid SM, stained with TMR-Star and examined using fluorescence microscopy. The cells were collected after 24 h of growth in liquid SM, stained with TMR-Star and examined by phase-contrast (PC) and fluorescence microscopy to monitor the SNAP production. The merged images show the overlap between the TMR-Star (red) and the autofluorescence (green) channels. The yellow arrows point the fluorescent signal. A schematic representation of the location pattern of the different proteins fused to SNAP Cd is shown on the right side of each panel. Among the cells with signal, the numbers represent the percentage of sporulating cells/spores showing the corresponding pattern of coat protein location. The data shown are representative of three independent experiments. For each strain, 100 cells were analysed. Scale bar, 1 µm.

Figure S6 :

 S6 Figure S6: Fluorescence localization of coat proteins fused to SNAP Cd . Microscopy analysis of the wild-type and cotL mutant cells carrying (A) CotD-and (B) CotE-SNAP Cd translational fusions. The cells were collected after 24 h of growth in liquid SM, stained with TMR-Star and examined by phase-contrast (PC) and fluorescence microscopy to monitor the SNAP production. The merged images show the overlap between the TMR-Star (red) and the autofluorescence (green) channels. The yellow arrows point the fluorescent signal. A schematic representation of the location pattern of the different proteins fused to SNAP Cd is shown on the right side of each panel. Among the cells with signal, the numbers represent the percentage of sporulating cells/ spores showing the corresponding pattern of coat protein location. The data shown are representative, of three independent experiments. One hundred cells were analysed for each strain. Scale bar, 1 µm.

cotL

  were amplified from genomic DNA using primer pairs CF126/CF127 (positions -243 to +438 from the translational start site) or CF189/CF127 (positions -75 to +438 from the translational start site). The resulting PCR products were inserted into pFT58 using Gibson Assembly yielding pDIA6617 and pDIA6663 and verified by sequencing. Plasmids were Results 96 introduced in E. coli HB101 (RP4) and transferred to C. difficile 630∆erm, sigE and sigK mutant strains by conjugation. To create other C-terminal SNAP-tag fusions, the promoter and coding regions of CD2629 (SpoIVA), CD1511 (CotB), CD1433 (CotE) and CD1067

  and anti-SNAP (1:1000) (NEB) were used. Anti-rabbit or anti-mouse secondary

  C. difficile, SpoIVA was recently shown to interact with SpoVM (Ribis et al. 2017) and SipL (Putnam et al. 2013). However, the lack of B. subtilis SafA and CotE homologs in C. difficile suggests that different factors are involved in the morphogenesis of the coat layer in this pathogen. Present in the coat layer and required for the assembly of proteins around the FS surface, CotL might play the role of morphogenetic proteins similar to B. subtilis SafA and CotE and to analyse it, we decided to search for potential CotL partners.

Figure 32 :

 32 Figure 32: CotL-SipL and CotL-SpoIVA pull down assays. Whole cell lysate and eluated fractions were tested by Western blotting using anti-CotL (A), anti-Histag (to detect SipL) (B) or anti-SpoIVA (C) antibodies. The indicated proteins were produced in E. coli upon induction with IPTG. Aliquots were removed, pelleted, and lysed to generate the whole-cell lysate sample. The lysate was applied into a Co 2+ affinity resin, eluted from the resin, and resolved on a 12% SDS-PAGE gel.

Figure 33 :

 33 Figure 33: CotL-CotB and CotL-CotE pull down assays. Whole cell lysate and eluated fractions were tested by Western blotting using anti-CotL (A), anti-CotB (B) or anti-CotE (C) antibodies. The indicated proteins were produced in E. coli upon induction with IPTG. Aliquots were removed, pelleted, and lysed to generate the whole-cell lysate sample. The lysate was applied into a Co 2+ affinity resin, eluted from the resin, and resolved on a 12% SDS-PAGE gel.

Figure 34 :

 34 Figure 34: Organization of the cotJ operons in B. subtilis and C. difficile. Open reading frames (arrows) and transcriptional start sites with promoters recognized by the sigma factors are indicated. The SpoIIID binding box in B. subtilis promoter is indicated in yellow. The percentage of amino acid identity between B. subtilis and C. difficile proteins is indicated in orange. In purple, is indicated the percentage of amino acid identity between C. difficile proteins from the two cotJ operons. σ K consensus elements were found upstream of CD2399 and CD0596. The positions of the -10 and -35 boxes are indicated in bolt.

Figure 35 :Figure 36 :

 3536 Figure 35: The CD2399 mutant is able to sporulate. A) Phase-contrast microscopy of sporulating cells of the wild-type and the CD2399 mutant strains after 24 h of growth in liquid SM. Yellow arrows mark mature phase-bright spores and red arrows indicate immature, phase-dark forespores. The numbers represent the percentage of sporulating cells. Scale bar, 2 µm. B) Counts (CFUs/mL) of total cells (black) and heat-resistant spores (grey) for the wild-type and the CD2399 mutant grown for 24 h in SM and plated on BHI with 0.1% TA. Data represent the average of four biological replicates.

Figure 37 :

 37 Figure 37: Morphology of the spores of the wild-type strain and the CD2399 mutant. A) Thin sections of the spores were analysed using TEM as described in the Materials and Methods. Ex, exosporium, Oc, outer coat, Ic, inner coat, Ct, cortex, Cr, core. Scale bars are shown in each figure. The numbers on the bottom of each figure represent the spore dimensions for each strain (length and width) based on the analysis of at least 8 individual spores. B) The thickness of the cortex layer and the exosporium plus the outer/inner coat layers of the spores from the two strains were measured. Error bars denote standard errors of the means of at least 8 individual spores. C) Size of the spore core of the wild-type and the CD2399 mutant. The length and width of the core was measured. Error bars denote the standard errors of the means of at least 8 individual spores.

Figure 38 :

 38 Figure 38: Protein profile of C. difficile spore coat/exosporium in the absence of CD2399. A) Coat/ exosporium extracts from purified spores of the wild-type, the CD2399 mutant and the complemented strains were loaded on acrylamide gel and stained with Coomassie brilliant blue. B) Western blot using antibodies raised against CdeM, CotE or SleC. Experiments were repeated at least two times with independent samples.

Figure 39 :

 39 Figure 39: Sensitivity of the wild-type and the CD2399 mutant spores to lysozyme, ethanol and heat. CFU/mL obtained of the wild-type and CD2399 mutant spores after plating onto BHI agar containing 0.1% TA. Data represent the average of the results of three independent experiments, and the error bars represent standard errors of the means.

  B. subtilis and likely controlled by σ K in C. difficile suggest an earlier production of the B. subtilis CotJ proteins. Consequently, B. subtilis CotJ-encoded products may be

  . It has been also noted that CotJC from B. subtilis carries a motif found in manganese catalases but enzymatic activity has never been demonstrated for this protein. The analysis of the localization of the C. difficile CotD protein similar to CotJC in the CD2399 mutant cells, and its assembly at the spore surface might be an interesting point to address in future work. So far, the role of CD2399 on the spore coat of C. difficile is not yet clear. Nonetheless, the presence of the paralogue CD0596 in the spore coat might suggest functional redundancy. Inactivation of the CD0596 gene in a CD2399 mutant background might bring new insights about the role of these two proteins in the assembly of the spore coat of C. difficile.

  enzymes have substrate-specificity to O 2 or NO but some FDPs have dual activity. Two genes encoding FDPs, CD1623 and CD1157, are present in C. difficile. On the other hand, Rbrs are composed by two types or iron sites: a N-terminal non-sulfur diiron center and a rubredoxinlike [Fe(SCys) 4 ] domain in the C-terminal part[START_REF] Demare | The structure of Desulfovibrio vulgaris rubrerythrin reveals a unique combination of rubredoxin-like FeS4 and ferritin-like diiron domains[END_REF], Kurtz 2006). Reverse Rbrs (revRbrs) are enzymes in which the position of the diiron site and the rubredoxin domains are reversed. The genome of C. difficile encodes two revRbrs, CD1474 and CD1524.

Results

Figure 40 :Figure 41 :

 4041 Figure 40: Transcriptional control of the CD1524 and the CD1623 genes. Promoter region of CD1524 (A) and CD1623 (B) genes. The mapped transcriptional start sites (+1, red) and the -10 and -35 promoter elements corresponding to the consensus for promoters recognized by σ B are indicated in bold (Soutourina et al, 2013). The possible RBS and the start codon of CD1524 and CD1623 are indicated in blue and orange, respectively. Microscopy analysis of C. difficile cells carrying P (CD1524) -SNAP Cd (C) and P (CD1623) -SNAP Cd (D) transcriptional fusions in the wild-type strain and in the sigB mutant. Cells were collected after 16 h of growth in BHI, stained with TMR and examined by fluorescence microscopy. The merged images show the overlap between the TMR-Star (red) and the autofluorescence (AF, green) channels. The numbers represent the percentage of cells with signal. Scale bar, 1 µm. These data are representative of 2 independent experiments.

  of the revRbrs and the FDPs in C. difficile vegetative cells I.1. Regulation of revRbrs (CD1474, CD1524) and FDPs (CD1623, CD1157) gene expression The genome-wide transcriptional start site (TSS) mapping study (Soutourina et al. 2013) identified a solely TSS for CD1474, CD1524 and CD1623 and two different potential TSS for CD1157. The promoter sequence upstream of the CD1474 TSS harbors a classical consensus sequence (WGWTT-N 13-17 -(G/T)GGTAWA) recognized by σ B in C. difficile (Kint et al. 2017). A σ B consensus sequence was also identified in the promoter region of the CD1524, CD1623 and CD1157 genes upstream of the TSSs (Figure 40A, B and Figure 41A, respectively). Moreover, upstream the second TSS of CD1157, we detected a promoter harboring a canonical -10 box (TATATT) of the σ A -dependent promoter and a less conserved -35 box (Figure 41A).

Figure 42 :Figure 43 :

 4243 Figure 42: Amino acid sequence of the FDP CD1157. The diiron center (35-224 aa) is represented in orange and the flavodoxin FMN domain (257-389 aa) in yellow. Amino acid residues that coordinate the catalytic iron atoms are in blue.

Results

Figure 44 :

 44 Figure 44: SDS-PAGE gel from the two revRbrs CD1524 and CD1474, and the FDP CD1157. Low-range molecular mass marker is indicated.

Figure

  Figure 45: UV-Visible spectra of CD1157 (A), CD1474 (B) and CD1524 (C). The line represents the full spectra of the oxidized proteins. In all the spectra, protein concentration was 30 µM and the experiments were performed in 50 mM Tris-HCl, pH 7.5 containing 18% glycerol.

Figure 46 :

 46 Figure 46: Anaerobic redox titration curves of the revRbrs CD1474 (A) and (B) CD1524. Panels correspond to the normalized intensities measured at 490 nm. In both titrations, protein concentration was 30 µM and the experiments were performed in 50 mM Tris-HCl, pH 7.5 containing 18% glycerol. The solid lines correspond to Nernst equations adjusted as described in the text, with the following reduction potentials: CD1474, +10 mV, CD1524, -10 mV.

Figure 49 :Figure 50 :Figure 51 :

 495051 Figure 49: Role of the two revRbrs in low O 2 tension tolerance. Serial dilutions of the wild-type strain, ∆CD1524, ∆CD1474 mutants, ∆CD1524-∆CD1474 double mutant, and the double mutant complemented with one or the one Rbr were spotted on TY taurocholate plates. Plates were then incubated 64 h either in anaerobic atmosphere (control) or in the presence of 0.1, 0.2 or 0.4 % O 2 .

Figure 52 :

 52 Figure52: Expression of the sigB gene and its target gene, CD1474, during sporulation. Temporal and cell type-specific expression of sigB (A) and of CD1474 (B) during sporulation. The cells were collected after 24 h of growth in liquid SM, stained with TMR-Star and the membrane dye MTG, and examined using fluorescence microscopy. The merged images showed the overlap between the TMR-Star (red) and the MTG (green) channels. The numbers represent the percentage of cells at the indicated stage that show expression of the reporter fusion. The panels are representative of the expression patterns observed for different stages of sporulation, ordered from early to late for the wildtype strains. Scale bar, 1 µm.

Figure 53 :

 53 Figure 53: Expression of the CD1524, CD1623 and CD0176 genes during sporulation. Schematic representation of the temporal and cell type-specific expression of CD1524 (A), CD1623 (B) and of CD0176 (C) during sporulation. The cells were collected after 24 h of growth in liquid SM, stained with TMR-Star and the membrane dye MTG, and examined using fluorescence microscopy. The numbers represent the percentage of cells at the indicated stage that show expression of the reporter fusion.

Figure 54 :Figure 55 :

 5455 Figure 54: Wild-type and sigB mutant spores. A) Morphology of the spores of the wild-type and sigB mutant. Thin sections of the spores were analysed using TEM as described in the Material and Methods. Ex, exosporium, Oc, outer coat, Ic, inner coat, Ct, cortex, Cr, core. Scale bars are shown in each figure. B) Coat/exosporium extracts from purified spores of the wild-type and the sigB mutant strains were loaded on acrylamide gel and stained with Coomassie brilliant blue.

  and T18. When T25 and T18 are separated, CyA is not active. However, when these fragments are fused with interacting proteins, the heterodimerization of these hybrid proteins allows a functional complementation and so the production of cyclic AMP (cAMP) leading to lacZ gene expression. The open reading frames of cotL, sipL, cotB, cotA, cotE and cdeC were cloned into plasmids containing T18 or T25 either in C-terminal or N-terminal position. The BTH101 E. coli strain was co-transformed by the different combinations of T18 and T25 fusion plasmids (TableS1). Co-transformation with the T25 and T18 expressing plasmids was used as control for the basal β-galactosidase activity. The βgalactosidase activity was measured after an overnight culture of co-transformants in the presence of 0.5 mM of IPTG (isopropyl-β-D-thiogalacto-pyranoside). 100 µl of overnight cultures was mixed with 500 µl of Z buffer (Na 2 HPO 4 0.06 M, NaH 2 PO 4 0.04 M, KCl 0.01M, MgSO 4 0.001 M, DTT 0.001 M and SDS 0.0037%) and 400 µl of M63 medium. After addition of 20 µl of chloroform, the tubes were mixed for 10 s. The tubes were pre-incubated 5 min at 28 °C and 200 µl ONPG (4 mg.ml -1 ) was added. When the yellow coloration appeared, 500 µl of Na 2 CO3 (1 M) was added to the tube to stop the reaction. After centrifugation to eliminate cells fragments, the supernatants were recovered to measure the OD 420nm . β-galactosidase activity was quantified as Miller unit [1000 x (OD420nm/(time of incubation (min) x volume of extract (ml) x OD 600nm of the overnight culture)].

  to +14 from the transcriptional start site) were amplified using primer pairs CF19/CF20, CF23/CF24 and CF21/CF22 to generate a 225 bp, a 221 bp and a 216 bp PCR product, respectively. The DNA fragment was then inserted into pFT47 between the EcoRI and XhoI sites to obtain pDIA6457, pDIA6459 and pDIA6458. To construct the CD0176transcriptional SNAP Cd fusion, the promoter region (-248 to -23 from the ATG start codon upstream of the first gene of the operon CD0174) was amplified using the primer pairs CF25/CF26 to generate a 231 bp PCR product. The DNA fragment was then inserted into pFT47 between the EcoRI and XhoI sites to obtain pDIA6460. The sequence of the inserts was checked by sequencing. The plasmids were introduced into E. coli HB101 (RP4) and then transferred by conjugation into C. difficile 630∆erm strain and the sigB mutant (Table

  antibodies raised againstCspC (1:2000), CspBA (1:2500), SleC (1:5000)(Adams et al. 2013), CdeM (1:40000) (A. O. Henriques), CdeC (1:1000), CotB (1:5000), CotE (1:1000), SipL (1:2500)(Putnam et al. 2013) and anti-SNAP (1:1000) (NEB) were used. Anti-rabbit or antimouse secondary antibodies conjugated to horseradish peroxidase (Sigma) were used at a 1:5000 dilution. Immunoblots were developed with enhanced chemiluminescence reagents (Thermo Scientific).

  potentials were investigated by EPR spectroscopy, using a Bruker EMX spectrometer equipped with an Oxford Instruments ESR-900 continuous flow helium cryostat, and a high sensitivity perpendicular mode rectangular cavity. The reduction potentials of the FMN from CD1157, and of the rubredoxin domain from the revRbr were determined by redox titrations monitored by visible absorption spectroscopy, in a Shimadzu UV-1603 spectrophotometer. Protein samples (30 µM in buffer C, 50 mM Tris-HCl, pH 7.5 and 18% glycerol) were anaerobically titrated, inside an anaerobic chamber (Coy Lab Products), by stepwise addition of a buffered sodium dithionite solution in the presence of a mixture of redox mediators (1 µM each): 1,2-naphthoquinone-4-sulfonic acid (E'° = +215 mV), trimethylhydroquinone (E'° = +115 mV), 1,4-naphthoquinone (E° = +60 mV), menadione (E'° = 0 mV), plumbagin (E'° = -40 mV), indigo trisulfonate (E'° = -70 mV), indigo disulfonate (E'° = -110 mV), 2-hydroxy-1,4-naphthoquinone (E'° = -152 mV) and anthraquinone-2-sulfonate (E'° = -225 mV). A combined Pt electrode (Ag/AgCl in 3.5 M KCl, as reference) was used and calibrated at 20°C against a saturated quinhydrone solution (pH 7). The reduction potentials are reported with the standard hydrogen electrode as a reference. All experimental data was manually adjusted to appropriate Nernst equations as described in the Results section: monoelectronic Nernst equations for the rubredoxin site, and Nernst equations for two consecutive monoelectronic transitions for the flavins, weighted according to the relative absorption intensities and concentrations of each cofactor.

  D(mrr-hsdRMS-mcrBC) f80lacZDM15 DlacX74 deoR, recA1 araD139 D(ara-leu)7697 galK rpsL(StrR) endA1 nupG NEB HB101 (RP4) supE44 aa14 galK2 lacY1 D(gpt-proA) 62 rpsL20 (Str R ) xyl-5 mtl-1 recA13 D(mcrC-mrr) hsdS B (r fhuA2 (lon) ompT gal (λ DE3) (dcm) ΔhsdS λ DE3= λ sBamHIo ΔEcoRI-B int::(lacI::PlacUV5::T7 gen1) i21 Δnin5 / E.coli strain for T7 expression, protease deficient B

Figure S1 :

 S1 Figure S1: Construction of C. difficile single and multi-mutants for the FDPs-and revRbrsencoding genes.For the CD1157 Clostron mutant, the verification of the L1.LtrB intron integration into the CD1157 gene was performed by using primers IMV844/IMV845 flanking the insertion site in CD1157. For the ACE mutants, the verification of the deletion of CD1623, CD1474 and CD1524 genes in the different backgrounds was performing using CF24/CF31, CF19/CF30 and CF23/CF39 respectively.

Figure 56 :

 56 Figure 56: Subcellular localization of the CotL protein fused to SNAP Cd . Microscopy analysis of C. difficile wild-type (A) and cdeM mutant (B) cells harboring P σE/σK(cotL) -cotL-SNAP Cd or P σK(cotL) -- cotL-SNAP Cd translational fusions. The cells were collected after 24 h of growth in liquid SM, stained with TMR-Star and examined by fluorescence microscopy to monitor the SNAP production. The merged images show the overlap between the TMR-Star (red) and the autofluorescence (green) channels. A schematic representation of the location pattern of the CotL-SNAP Cd protein is shown on the right side ordered from early to late stages. Among the cells with signal, the numbers represent the percentage of sporulating cells/spores showing the corresponding pattern of coat protein location. Scale bar, 1 µm.

Figure 57 :

 57 Figure 57: Model of the morphogenetic proteins of the outermost layers of the C. difficile spore. A) Preliminary model of the localization of the spore coat (SpoVM, SpoIVA, SipL and CotL) and exosporium (CdeC and CdeM) morphogenetic proteins. The hexagon represents the LysM domain. B) Proteins interaction network. Confirmed protein interactions are represented by a double arrow. Double dotted arrows represent positive interactions determined by BACTH. One-way orange arrows indicate proteins which assembly or recruitment at the FS is dependent on CotL. One-way pink arrows indicate proteins which abundance in the spore coat/exosporium is dependent on CdeC or CdeM.

  like in B. subtilis. Based on our results including the early and late expression of cotL gene during sporulation, the morphology of the cotL mutant spores and the requirement of CotL for the localization of other spore coat proteins, we propose that CotL may be located in the inner coat layer. In B. subtilis, SpoIVA and SpoVID interact with CotE and SafA. Since no homologs of these proteins are present in C. difficile, further research is needed to identify additional morphogenetic spore coat proteins recruited by SpoIVA, SipL, SpoVM and/or CotL.

Figure 58 :

 58 Figure 58: Model of the signal transduction leading to the activation of σ B in C. difficile. Partner switching mechanism of σ B activation is conserved in C. difficile. During stationary growth phase or after stress exposure, the RsbZ phosphatase dephosphorylate RsbV, which interacts with RsbW, allowing σ B release, its association with the RNA polymerase and transcription of its targets genes. Stress signals triggering phosphatase activation in C. difficile are not know but exposure to CCCP and entry into stationary phase activate σ B activity.

σ

  B activity. In other Firmicutes, exposure to environmental stresses typically induces the activation of σ B (Hecker et al. 2007). In B. subtilis and L. monocytogenes, these signals are detected via the stressosome by an unknown mechanism (Pane-Farre et al. 2017). In B. cereus, the stressosome is absent, and the activation of σ B by environmental signals appears to be mediated by the kinase RsbK (de Been et al. 2010). The absence of homologues of the stressosome components or RsbK in C. difficile might explain the absence of induction of σ B activity in response to the various environmental stresses tested. However, we cannot exclude the possibility that the conditions used to test σ B induction are not drastic enough to promote the activation of this sigma factor in C. difficile.

  data suggest that C. difficile does not face a nutrient-deficient environment during gut colonization. On the other hand, most of the C. difficile proteins involved in energy metabolism and maintenance of oxidation-reduction potential contain cysteines and Fe-S clusters. These molecules are extremely sensitive to oxidation and are the main targets of the oxidizing agents found during the infectious cycle such as O 2 and/or ROS, NO or RNS produced during inflammation(Py et al. 2011). Therefore, it is possible that the activation of σ B through energy stress is due to an indirect effect of the oxidative environment encountered by C. difficile during its infectious cycle including the inactivation of enzymes involved in the central metabolism of the bacterium. Further studies will be performed to decipher the role of the energy metabolism and the mechanisms involved on σ B activation in C. difficile.

Figure 59 :

 59 Figure 59: Heterogeneity of C. difficile gene expression. At stationary phase, the C. difficile bacterial population likely specialize into distinct subpopulations responsible for sporulation, σ Bmediated stress response and toxin production. The two last subpopulations can survive as vegetative cells or eventually sporulate allowing the activations of σ B or the toxin production in sporulating cells.

  study of Ramson et al, and our data indicate that sporulation, toxin production and σ Bdependent response are not exclusive processes. Toxin gene expression and sporulation can occur in the same cell and σ B is active in sporulating cells, as it will be discussed more in detail below. These suggest that under stressful conditions, a sub-population of cells might directly initiate sporulation, while other sub-populations develop mechanisms of stress response mediated by σ B or use their energetic sources to produce the toxins (Figure59). In the presence of a currently unknown stimulus, these two last groups of cells can survive as vegetative cells or eventually might enter into sporulation. Whether σ B activity and toxin production occur in the same cell is not yet known. In order to address this point, an interesting strategy would be to use different fluorescent reporter genes fused to promoters involved in stress response and toxin synthesis to study gene expression within a bacterial population by fluorescence microscopy.

Figure 61 :

 61 Figure 61: Model of the O 2 and ROS detoxification pathway in C. difficile. Scheme of the NAD(P)H-dependent O 2 , O 2 -and H 2 O 2 detoxification complex. Structural domains are color coded. NROR (NADH rubredoxin oxidoreductase), OR (oxidoreductase), Rd (Rubredoxin domain), FMN (flavodoxin domain) and Fe-Fe (diiron center). Arrows indicate eletron transference. Candidates for the OR may be CD0176, CD2540 and CD2709. The CD0827 desulfoferrodoxin is a candidate for the reduction of the O 2 -.

Figure 62 :

 62 Figure62: Model of the negative control of σ B on C. difficile sporulation. The initiation of sporulation is dependent on σ H , Spo0A and its phosphorylation state. σ B negatively and positively controls the expression of the CD1579 and the CD1492 genes, respectively. σ B negatively controls sporulation likely through regulation of the Spo0A phosphorylation. Green and red arrows represent positive or negative control respectively. A mechanism of phosphatase activity for CD1492 have not been characterized.

  regulation factors controlling the σ B regulon during sporulation. The detailed mechanisms controlling the expression and activation of σ B during sporulation will be addressed in future experiments. IV.3. The hypothetical protective role of the σ B -target genes IV.3.a. Spore resistance

Figure 63 :

 63 Figure 63: Phase-contrast microscopy of purified spores of the wild-type and the ΔCD1474ΔCD1524 double mutant strains. Green arrows mark mature phase-bright spores and red arrows indicate immature, phase-dark spores.

Figure 64 :

 64 Figure 64: Model for the role of σ B and sporulation during colonization of the gastrointestinal tract by C. difficile. After ingestion,C. difficile spores will germinate in the intestine. Once in the colon, vegetative cells are exposed to various stresses such as the increasing O 2 tension in the proximity of the epithelial cells. In addition, the host produces oxidizing compounds such as ROS/NO and RNS via the inflammatory process. The activation of σ B and the synthesis of the FDPs and revRbrs proteins will allow bacteria to survive and proliferate in the colon. In the presence of a yet not known stimulus, C. difficile sporulates and in a sub-population of sporulating cells, σ B is active. In the MC compartment, the spore coat protein CotL, which encoding gene is transcribed by both σ E and σ K , localizes first as a cap in the MC proximal pole and later around the spore. CotL is responsible for the encasement of other coat proteins around the FS, playing a key role in the morphogenesis of the C. difficile spore coat.

  germination. Then, C. difficile vegetative cells multiply and colonise the intestinal tract. The toxigenic strains of C. difficile produce two toxins, TcdA and TcdB, which are the major virulence factors required for the development of symptoms. These toxins cause the alteration of actin cytoskeleton in intestinal epithelial cells that leads to a major neutrophil recruitment favouring an intensive local inflammation process(Peniche et al. 2013, Jose and Madan 2016). Although toxins are regarded as primary virulence factors, the establishment of C. difficile in its colonic niche is also dependent on additional virulence factors such as the adhesin Cwp66, the flagellum, the fibronectin-binding protein FbpA, and the surface layer precursor protein SlpA(Janoir 2016).In the gut, C. difficile vegetative cells face different stresses including low physiologic oxygen (O 2 ) tensions(Espey 2013, Rivera-Chavez et al. 2017), pH variations, antibiotics and hyperosmolarity. Bacteria are also exposed to bile acids and cationic antimicrobial peptides (CAMPs) produced by the host and/or by the microbiota(Abt et al. 2016b). Moreover, the inflammation process triggered by several C. difficile factors (toxins, S-layer proteins and flagellin) leads to the production by the host immune system of reactive oxygen species (ROS), nitric oxide (NO) and reactive nitrogen species (RNS)(Abt et al. 2016b, Jose and Madan 2016). Thus, to adapt and colonise this hostile environment, C. difficile must induce protection, detoxification and repair systems. We have recently shown that the alternative sigma factor of the general stress response, σ B (

Figure 1 :

 1 Figure 1: Genetic organisation of the CD0007-CD0011 locus. Organisation of sigB operon.Gene function is indicated below each gene. TSS mapping experiment indicated the presence of a σ A -dependent promoter upstream of CD0007(Soutourina et al, 2013). The -35 and -10 boxes as well as the TSS are indicated in bold. PCR were both realised on RNAs without reverse transcription (lane 1, 3, 6 and 8) and on cDNA synthesised from the same RNAs by reverse transcription using IMV389 primer (lane 2, 4, 7 and 9). PCR was then performed with primers annealing in CD0007 and CD0008 (IMV774/OS548, lane 1 and 2), in CD0008 and CD0009 (OS547/IMV777, lane 2 and 3), in CD0009 and CD0010 (IMV776/IMV779, lane 6 and 7) and in CD0010 and CD0011 (IMV778/IMV389, lane 8 and 9). Lane 5: Ladder.

Figure 2 :

 2 Figure2: sigB overexpression is toxic for C. difficile. Growth curves of 630∆erm strain containing either pDIA6103 (A) or P tet -sigB (B) in TY in presence of different concentrations of aTc. These data are the mean of 4 independent experiments. C) Fold change of the expression of rsbV, rsbW and sigB genes and of some σ B target genes between 630∆erm + P tet -sigB and 630∆erm + pDIA6103 without aTc (black), with 25 ng.ml -1 (dark gray) or 200 ng.ml -1 of aTc (light gray). These results are the means of 2 independent experiments.

Figure 3 :

 3 Figure 3: Conservation of the partner switching mechanism in C. difficile. Serial dilutions of the different strains were spotted on TY plates containing 25 ng.ml -1 of aTc. Plates were incubated 24 h at 37°C for NO tests (A) or 64 h in presence of 0.4 % of O 2 for low O 2 tolerance assay (B).Growth controls were performed with plates supplemented with NaOH or plates incubated in anaerobiosis atmosphere for NO tests and low O 2 tolerance assay, respectively. These pictures are representative of 3 independent experiments. C) Effect of rsbV deletion or rsbW overexpression on the expression of sigB operon genes and σ B target genes at the onset of stationary phase. Fold change of the differential expression of the genes belonging to the sigB operon and σ B target genes involved in oxidative and nitrosative stress management in 630∆erm ∆rsbV + pDIA6103 (black), 630∆erm ∆rsbV + P tet -rsbV (light gray) and 630∆erm + P tet -rsbW (dark gray) compared to 630∆erm + pDIA6103. These results are the mean with the standard deviation of 4 independent experiments.

Table 2 :

 2 Interaction of the different proteins involved in the σ B activation pathway in bacterial adenylate cyclase two hybrids. Values shown are the average ß-galactosidase activities (in Miller unit) of the indicated pair-wise fusions. Each value are the means and the standard deviation of 4 or 5 independent experiments.

  . To identify such PP2C phosphatases in C. difficile, we looked for proteins similar to RsbP and RsbU, the two phosphatases involved in RsbV dephosphorylation in B. subtilis. Two PP2C phosphatases are encoded in C. difficile genome: SpoIIE (CD3490), involved in the sporulation process (E. Cuenot and I. Martin-Verstraete, unpublished results), and CD2685. The last 200 amino acid residues of CD2685 share 27 % and 25 % identity with RsbP and RsbU, respectively. The C-terminal region of CD2685 also shares similarities with other PP2C phosphatases involved in σ B activation in pathogenic firmicutes such as L. monocytogenes, B. cereus and S. aureus. No trans-membrane helices or signal peptides are predicted for CD2685. CD2685 protein contains three domains: i) the well characterised serine/threonine PP2C phosphatase domain in the C-terminal part (amino acid 369 to 588), ii) a putative coiled coil domain (amino acid 335 to 360) and iii) a putative sensory GAF domain (amino acid 31 to 183). GAF domains are found in several proteins including cGMP-specific phosphodiesterase, adenylyl-cyclase, guanylyl-cyclase or phytochrome (Aravind and Ponting 1997). Based on these in silico analysis, we hypothesised the possible involvement of CD2685 in the activation of σ B .

Figure 4 :

 4 Figure 4: CD2685/RsbZ phosphatase is responsible for σ B activation in C. difficile. A)Effect of rsbZ deletion of or its phosphatase domain (rsbZ∆pho) on the expression of sigB operon genes and σ B target genes at the onset of the stationary phase. Fold change of the differential expression of the genes belonging to sigB operon, rsbZ operon and of σ B target genes involved in oxidative and nitrosative stress management in 630∆erm ∆rsbZ + pDIA6103 (black), 630∆erm ∆rsbZ + P tet -rsbZ (light gray) and 630∆erm ∆rsbZ + P tet -rsbZ∆pho (dark gray) compared to 630∆erm + pDIA6103. Low O 2 tolerance (B) and NO stress assays (C). Serial dilutions of the different strains were spotted on TY plates containing 25 ng.ml -1 of aTc. Plates were incubated 64 h in presence of 0.4 % of O 2 (B) or 24 h at 37°C for NO tests (C). Growth controls were performed in anaerobiosis atmosphere or in presence of NaOH for low O 2 tolerance assays and NO tests, respectively. D) RsbV phosphorylation state in wild-type and ∆rsbZ backgrounds. Cellular extracts were run on 12 % acrylamide gel containing 50 µM of Phos-Tag™ compounds allowing the separation of phosphorylated and non-phosphorylated proteins. Western blot was performed with an antibody raised against the HA tag. Loading control was realized using a 12 % acrylamide gel followed by Coomassie staining. qRT-PCR results are the mean with the standard deviation of 4 independent experiments and pictures are representative of 4 independent experiments.

Figure 5 :

 5 Figure5: The essential protein CD2684 is involved in σ B signalling pathway. A) Construction of conditional CD2684 mutant. The entire P tet element from pRPF185(Fagan and Fairweather 2011) was inserted into the chromosome between rsbZ and CD2684 allowing CD2684 expression of under the control of P tet and its induction following aTc addition. B) CD2684 is essential in C. difficile. Growth phenotype of 630∆erm (left part) or sigB mutant (right part) carrying a chromosomal P tet CD2684 copy on TY plates without aTc or with 10 and 25 ng.ml -1 of aTc after 24 h at 37°C. C) Growth of 630∆erm chromosomal P tet CD2684 in TY with different aTc concentrations. D) Effect of CD2684 expression shut down on the expression of sigB operon and σ B target genes. Fold change of the differential expression of sigB, of genes of rsbZ operon and of different σ B target genes encoding proteins involved in oxidative and nitrosative stress management between 630∆erm chromosomal P tet CD2684 grown without aTc or with 10 ng.ml -1 of aTc. The graph shows the expression fold change after 4 h (black) or 10 h (light gray) of growth. Pictures are representative of 3 independent experiments and growth results represent the mean of 3 independent experiments. qRT-PCR results are the mean with the standard deviation of 3 independent experiments.

Figure 7 :

 7 Figure 7: Model of σ B signalling pathway in C. difficile. Established and predictive signalling pathway for the control of σ B activity in C. difficile. Environemental or energeticstresses not yet identified are indicated by the "flash" symbol. Locations of potential stimulus perception are indicated by "explosion" symbols. The CD2684 essential iron-sulphur flavoprotein might be involved in stress detection or be involved in cellular process that results in RsbZ phosphatase activation. This activation leads to RsbV dephosphorylation that enables its interaction with the RsbW anti-sigma factor. This protein-protein interaction results in σ B release, its association with RNA-polymerase core enzyme and the transcription initiation of its target genes.

  C. difficile. In B. subtilis and B. cereus, the input N-terminal domain of the phosphatases, RsbP and RsbY, is a PAS (Per-Arnt-Sim) or a REC (signal receiver domain)

Figure S1 :

 S1 Figure S1: Overexpression of genes involved in the partner switching mechanism. Growth curves of the 630∆erm + P tet -rsbV, 630∆erm + P tet -rsbW, 630∆erm ∆rsbZ + P tet -rsbZ, 630∆erm + P tet -rsbZ and 630∆erm ∆rsbZ + P tet -rsbZ∆pho in presence of different concentration of aTc. These data are the mean of 4 independent experiments.

Figure S2 :

 S2 Figure S2: Western Blot. A) Production of σ B HA-tagged protein from overnight culture. (1) 630∆erm + pB-sigB, (2) 630∆erm + pB-sigB-HA, (3) sigB + pB-sigB-HA, (4) 630∆erm ∆rsbV + pB-sigB-HA and (5) 630∆erm ∆rsbZ + pB-sigB-HA. B) Verification of the production of RsbZ∆pho protein.(1) 630∆erm ∆rsbZ + P tet -rsbZ, (2) 630∆erm ∆rsbZ + P tet -rsbZ-HA and (3) 630∆erm ∆rsbZ + P tet -rsbZ∆pho-HA. Western blot using an antibody raised against the HA tag. Loading control was realized using a 12 % acrylamide gel followed by Coomassie staining.

Figure S4 :

 S4 Figure S4: CD0007 and CD0008 are not involved in σ B signalling pathway. Low O 2 tolerance (A) and NO stress assays (B). Serial dilutions of the different strains were spotted on TY plates. Plates were incubated 64 h in presence of 0.4 % of O 2 for low O 2 tolerance assay (A) or 24 h at 37°C for NO tests (B). Growth controls were performed with plates incubated in anaerobiosis atmosphere or supplemented with NaOH for low O 2 tolerance assays and NO tests, respectively. C) Effect of CD0007, CD0008 or CD0007-08 deletion on the expression of sigB operon genes and σ B target genes at the onset of the stationary phase. Fold change of the differential expression of genes belonging to sigB operon, to rsbZ operon and of σ B target genes involved in oxidative and nitrosative stress management in 630∆erm ∆CD0007 (black), 630∆erm ∆CD0008 (light gray) and 630∆erm ∆CD0007-08 (dark gray) compared to 630∆erm. Pictures are representative results of 3 independent experiments and qRT-PCR results are the mean with the standard deviation of 3 independent experiments.

  pDIA6637 and pDIA6579, carrying sigB, rsbZ or rsbZ∆pho were inverted PCR amplified using primers NK188/NK189, NK192/193 and NK194/195, respectively. PCR fragments were then digested by DpnI to remove the plasmid templates, phosphorylated by T4 Polynucleotide Kinase and ligated by T4 Ligase to recircularize the plasmid. The resulting plasmids were verified by sequencing, introduced into HB101/RP4 E. coli strain and then transferred into C. difficile strains by conjugation. Transconjugants were selected on BHI plates supplemented with Tm and C. difficile selective supplement.

  β-galactosidase activity was quantified as Miller unit [1000*(OD 420nm /(time of incubation (min)*volume of extract (ml)*OD 600nm of the culture)].

  CDIP1117630∆erm sigB::erm + pDIA6103-P tet -rsbV HA C-ter pDIA6639 → CDIP546 CDIP1118 630∆erm ∆rsbV + pDIA6103-P tet -rsbV HA C-ter pDIA6639 → CDIP713 CDIP1119 630∆erm ∆rsbZ + pDIA6103-P tet -rsbV HA C

  TAG ATA CAG AGT CAT TA qRT-PCR CD1065 CF131 TGC CAT CTA GAT TCT

Oxygen and Reactive Oxygen Species detoxification I. Cell defense mechanisms against these compounds present in bacteria I.1. Catalases and superoxide dismutases

	O 2	. OH + H 2 O Chapter 3: Mechanisms of cell defense against oxidative stress have been extensively studied in O 2 .-H 2 O 2	H 2 O
	aerobic organisms. In this case, oxidative stress is a by-product of the aerobic lifestyle (Imlay
	2003). Microorganisms typically deal with oxidative stress using two major enzymes:
	superoxide dismutases (SODs) and catalases. SODs catalyze the dismutation of the
	superoxide (O 2	-) into O 2 and H 2 O 2 , a reaction requiring one proton per O 2	-reacted, but no
	external reductant. Then, H 2 O 2 is degraded to O 2 and water by catalases (Figure

  Production of O 2 by the SOD/catalases is obviously a disadvantage for anaerobes.

Nevertheless, although strictly anaerobic, microorganisms such as C. acetobutylicum and Clostridium perfringens are considered to be aerotolerant anaerobes. C. acetobutylicum survives short periods of aeration (O'Brien and Morris 1971) while C. perfringens vegetative and stationary-phase cells can survive in a growth-arrested state in the presence of O 2 and/or low concentrations of O 2 --or hydroxyl-radical-generating compounds

  . Rd increases the NADH consumption rate by doing the electron-shuttle between NROR and Rbrs. In C. acetobutylicum, which does not have σ B , the O 2 and H 2 O 2 detoxification pathways are controlled by the negative regulator PerR, that senses H 2 O 2 as a signal of oxygenation

  The host NADPH oxidase NOX2 complex activated by both phagocytosis of bacteria and by TcdB produce ROS such as O 2 -and H 2 O 2 in the phagosome (Decoursey and Ligeti 2005). In addition, NO produced by nitric oxide synthases, mainly by iNOS, can react with O 2 -and form peroxynitrite (ONOO -), nitrogen dioxide and other toxic RNS. Both ROS and RNS can

	react with multiple cellular components being deleterious to nearly all biomolecules. By
	mainly oxidizing Fe-S clusters, O 2	-can result in the inhibition of enzymatic activity and a

  ). Nevertheless, the response of C. difficile to atmospheric O 2 and H 2 O 2 exposure suggested that this pathogen has mechanisms to tolerate oxidizing conditions. However, the mechanisms used by C. difficile to resist exposure to H 2 O 2 , air or O 2

and to detoxify these compounds are not yet clearly identified. It was demonstrated that σ B plays a crucial role in this tolerance

(Edwards et al. 2016a, Kint et al. 2017)

. Moreover, the inactivation of the gluD gene encoding the glutamate dehydrogenase GluD (CD0179) responsible for the degradation of glutamate into α-ketoglutarate and NH3, increases the sensitivity of C. difficile to H 2 O 2

  ). C. difficile very likely encodes other functions allowing the bacterium to tolerate and survive in the presence of low O 2 tensions and during oxidative stress. In 2008, Emerson et al performed a large-scale transcriptomic study that provided some insights into the response of this

bacterium to air stress. The oxidative shock created by the exposure to air increased the expression of 98 genes in C. difficile, including several genes belonging to the σ B regulon involved in oxido-reduction and electron transfers

Chapter 4: Sporulation and Germination I. General stress response and sporulation: two survival mechanisms in C. difficile

  B , but not by σ H . On the contrary, σ H represses the expression of C. difficile toxin genes, whereas σ B does not play any role in toxinogenesis.

	Bacterial cell entry into stationary phase implies changes in metabolism and physiology. σ B ,
	, the presence in the C. difficile genome of genes encoding proteins the known sigma factor of general stress response, participates in the reprogramming of C.
	sharing similarities with revRbrs and FDPs from C. acetobutylicum suggests the presence of difficile cellular metabolism in stressful environments or at the onset of the stationary phase
	O 2 and ROS alternative detoxification pathways in this bacterium. The C. difficile genome (Kint et al. 2017). When cells enter stationary phase (~10 h), it was shown that σ B controls the
	encodes a desulfoferrodoxin (CD0827), two revRbrs (CD1524 and CD1474), two Rbrs expression of approximately 25% of C. difficile genes (Kint et al. 2017). Moreover, the sigma
	(CD0825 and CD2845), two FDPs proteins (CD1157 and CD1623) and several OR (CD0176, factor σ H , which plays a crucial role in C. difficile during transition phase, controls the
	CD1623 and others). With the exception of the two Rbrs, all of these proteins are encoded by expression of several genes involved in metabolism, motility and sporulation (Saujet et al.
	genes controlled by σ B and a σ B -dependent promoter is mapped or identified in silico 2011). Thus, σ B together with σ H , control around 49% of the genes induced at the onset of the
	upstream of all these genes (Kint et al. 2017). These data were in agreement with the control stationary phase and are the major sigma factors in decision making during post-exponential
	by σ B on the resistance to ROS and the tolerance to low O 2 tensions (Kint et al. 2017). A very growth (Saujet et al. 2011, Kint et al. 2017). Interestingly, genes involved in oxidative-and
	recent study has functionally characterized CD1623 as an NADH:O 2 reductase that might nitrosative-stress responses are controlled by σ
	contribute to the survival of C. difficile in the gut (Folgosa et al. 2018b) (Figure 15). This
	protein belongs to the Class F of FDPs and contains a diiron center and an FMN in the core
	domain, a FeCys 4 center at a short spaced rubredoxin domain (as in classical Rbrs) and a
	NROR domain (binding a FAD) at the C-terminal (Figure 15). It is worth noting that CD1623
	carries into a single polypeptide a complete electron transfer chain that enables coupling
	directly O 2 reduction to the oxidation of the electron donor NAD(P)H. This enzyme has a
	clear preference to O 2 as a substrate. Except for CD1623, the role of the other C. difficile
	proteins with a possible function in O 2 and ROS detoxification has not been studied. In the
	framework of my PhD goals, I studied the O 2 and ROS detoxification pathways in C. difficile,
	focusing mainly on the enzymatic characterization of the other FDP, CD1157, and the two
	revRbrs CD1474 and CD1524, and their role for this anaerobic pathogen.

sigK Pσ E , SpoIIID SpoIVCA skin Pσ E , SpoIIID Pro-σ K σ K SpoIVFB sigK Pσ E Pσ K SpoIIID σ K CD1231 CD1234 Pσ A skin Pσ E , SpoIIID skin Bs skin Cd Figure 22: Regulation of σ K activity in B. subtilis and in C. difficile. A prophage-like skin element must excise from genomic DNA prior to form a functional sigK gene. A) In B. subtilis, skin excision

  is meditated by the recombinase SpoIVCA. Then, σ E and the SpoIIID regulator control sigK expression. σ K is produced as an inactive pro-σ K that undergoes proteolytic activation via the SpoIVFB protease. B) In C. difficile, the recombination directionality factor, CD1234, produced in the MC forms a complex with the recombinase CD1231 to catalyze skin excision. The rearrangement of the sigK gene leads to the production of an intact and functional sigma factor. Adapted from Fimlaid and Shen 2015.

	A	B
	morphogenetic proteins) (McKenney and Eichenberger 2012). σ E drives the activation of σ G
	B. subtilis	C. difficile
	Additionally, σ E controls the expression of the the coat morphogenetic proteins spoVM,
	spoIVA, spoVID, safA and cotE (see more in detail Chapter 5, section III.2.a. Coat

  . In addition, the regulation and activation of the four sporulation σ factors display different patterns among clostridia. For instance, σ F and σ E are active prior to asymmetric division in C. acetobutylicum and C. perfringens. In C. acetobutylicum, the σ F regulon is quite different from that of B.

Pro-σ E SpoIIGA SpoIIID MC FS SpoIIR SpoIIIA-H SpoIIQ SpoVT OFM IFM σ G regulon sspA, sspB, , spoVA, pdaA σ E regulon

	cwlD, spoIID,
	spoVA, cotB	σ F
		regulon
		gpr,
		spoIIP,
		spoIIQ
	σ K regulon
	cotA, cotE, cotD,
	bclA1, sleC

Pσ A spoIIGA sigE Spo0A-P sigG Pσ F/ σ G Pσ H spoIIAA spoIIAB Spo0A-P sigF Figure 24: Model of the regulatory network controlling sporulation in C. difficile. Temporal

	progression of sporulation is shown from top to bottom. Boxes represent the predivisional cell (PDC),
	mother cell (MC) and forespore (FS), the two cell compartments formed after asymmetric division.
	Membranes separating the FS and the MC are indicated (OFM: outer forespore membrane; IFM:
	inner forespore membrane). Sigma factors are colored and bold, the precursor protein of σ E is
	indicated as pro-σ E . Transcriptional regulators are shown in purple. Proteins associated with the
	membranes or located into the intermembrane space are in square. Black solid arrows indicate
	activation at the transcriptional level. Red arrows indicate the intercellular signal pathways required
	for sigma factor activity. Broken arrows with a question mark represent mechanisms that are not yet
	fully understood. Adapted from Saujet et al., 2013.

?

?

? Pσ A /other? σ E σ F σ G σ K SpoIIE SpoIIAA SpoIIAA σ F SpoIIAB P difficile

  , σ K is dispensable for cortex development and controls coat polymerization, similar to B. subtilis. Even if the main functions and periods of activity of the sporulation specific sigma factors are conserved in C. difficile, a less tight communication between the FS and the MC compartment and a weaker connection between morphogenesis and expression are observed

Article: CotL, a new morphogenetic spore coat protein of Clostridium difficile

  . difficile spores play a central role in the infectious cycle, contributing to transmission, infection and persistence(Lawley et al. 2010, Deakin et al. 2012). The spore surface layers i.e., the coat and exosporium, enable spore resistance to enzymatic degradation and chemicals present in the host gastrointestinal tract[START_REF] Riesenman | Role of the spore coat layers in Bacillus subtilis spore resistance to hydrogen peroxide, artificial UV-C, UV-B, and solar UV radiation[END_REF]

I.

C

  al. 2013, Diaz-Gonzalez et al. 2015). The exosporium is absent in B. subtilis, and only 25% of the coat proteins have homologs in C. difficile(Henriques and Moran 2007, Permpoonpattana et al. 2011b, Permpoonpattana et al. 2013). The spore surface of the bacilli is primarily composed of proteins produced in the MC under the control of first σ E and later σ SpoIVA recruits the morphogenetic protein from the inner coat, SafA, while both SpoIVA and SpoVID are required for the recruitment of CotE, an outer coat morphogenetic protein(de Francesco et al. 2012, McKenney et al. 2013).

K (Henriques and Moran 2007, McKenney et al. 2013

). In addition to transcriptional regulation, coat assembly depends on the ordered synthesis and recruitment of proteins that are encased at the FS surface forming a basement layer, an inner coat and an outer coat. In B. subtilis, the formation of these three layers is governed by morphogenetic proteins, such as SpoVM, SpoIVA, SpoVID,

SafA and CotE (McKenney et al. 2013)

. Homologs for SpoIVA and SpoVM are present in C. difficile

(Galperin et al. 2012)

. The SpoIVA of C. difficile is a morphogenetic protein essential for coat localization around the FS at the early development stages of sporulation

(Putnam et al. 2013

). The C. difficile SpoIVA directly interacts with SpoVM even if SpoVM is not essential to encase SpoIVA in the FS. SpoVM is also dispensable for spore formation

(Ribis et al. 2017

) in contrast to that observed in B. subtilis. SipL has been identified as a morphogenetic protein specific to C. difficile. SipL, together with SpoIVA, interacts and drives coat assembly around the C. difficile FS

(Putnam et al. 2013)

. SipL is likely to be a functional homologue of SpoVID of Bacillus sp., a protein that helps in the transition of the spore coat from the mother cell proximal (MCP) pole to full encasement

(Wang et al. 2009a, McKenney and Eichenberger 2012)

. In B. subtilis,

Table 1 . Spore surface proteins less abundant in the CD1065 mutant compared to the wild-type strain.

 1 which lacks the inner membrane germinant receptors from the Ger family known to sense amino acids and sugars, uses a CspC pseudoprotease located in the coat extractible fraction to sense bile acid germinants, such as TA(Sorg and Sonenshein 2008, Paredes-Sabja et al. 2011, Francis et al. 2013a). When CspC senses the germinant, it activates the serine protease CspB that will cleave Pro-SleC in SleC, the cortex hydrolase. Cortex hydrolysis triggers Ca-DPA release

	Protein (Gene)	MW	Function	Fold-	p-value	Know/predicted
		[kDa]		change (WT/CD1065)	(LIMMA t-test)	σ factor needed for expression
	CspBA (CD2247)	125	Serine protease processes Pro-SleC	2	1,7E-05	σ E
			into SleC
	SleC (CD0551)		Spore cortex-lytic enzyme	10	8,5E-08	σ K
	SpoIVA (CD2629)		Morphogenetic coat protein	3	6,2E-08	σ E
	SipL (CD3567)		SpoIVA-Interacting protein	3	6,5E-08	σ E
	CotA (CD1613)		Coat protein	2	6,4E-06	σ K
	CotB (CD1511)		Outer coat layer protein	45	1,1E-10	σ E
	CotE (CD1433)		Coat protein Peroxiredoxin/chitinase	3	1,1E-06	σ K
	CotG (CD1567)		Coat protein Manganese catalase	3	3,7E-06	σ G
	CD0596	9	Hypothetical protein	7	1,3E-09	σ K
	CotF (CD0597)		Spore coat protein	8	9,9E-09	σ K
	CotCB (CD0598)		Coat protein manganese catalase	3	1,1E-07	σ K
	CD2399	8	Hypothetical protein (CotJA	9	2,9E-06	σ K
			superfamily)
	CD2400		Spore coat assembly protein CotJB2	6	3,1E-06	σ K
	CotD (CD2401)		Coat protein manganese catalase	2	6,6E-06	σ K
	Rbr (CD2845)		Rubrerythrin	4	3,6E-06	σ G
	SodA (CD1631)		Superoxide dismutase	44	2,0E-10	σ G
	CD3580		Hypothetical protein	7	1,3E-09	σ K
	CdeC (CD1067)		Cysteine-rich exosporium	3	1,5E-05	σ K
			morphogenetic protein
	CdeM (CD1581)		Conserved hypothetical protein	3	6,7E-07	σ K
	BclA1 (CD0332)		Exosporium glycoprotein	4	3,1E-06	σ K

Results

84

spores are defective in the earliest stages of germination.

The germination-signalling pathway of C. difficile is markedly different from that of the other spore-forming bacteria

(Bhattacharjee et al. 2016)

. C. difficile,

  The anti-CotL antibody used in this study was raised in mice by Agro-Bio. CotL-His 6 was purified on Co 2+ affinity resin from E. coli strain 832. Cultures were grown and protein expression was induced in Terrific broth (TB) medium

	(Thermo Scientific) for 16h at 37°C. E. coli cells were harvested, resuspended in lysis buffer
	buffer A (50 mM NaH 2 PO 4 , 300 mM NaCl, 10 mM imidazole) containing 1X Bug buster

(Novagen), 40 µg/mL Dnase and 50 µg/mL lysozyme) and lysed at RT for 40 min. Cell extracts were centrifuged, the supernatant was passed through a 0.2 µM filter and used to affinity purify CotL-His 6 . Elution was performed using buffer B (50 mM NaH 2 PO 4 , 300 mM NaCl, 300 mM imidazole).

Table 1 : Determination of the interaction of CotL with other different spore coat proteins using bacterial adenylate cyclase two hybrid (BACTH).

 1

	CotL + SipL	pUT18C		pUT18
			CotL	SipL	CotL	SipL	-
		CotL		1136±758		960±788
	pKT25	SipL	742±425	1526±475
		CotL		767±394		212±78
	pKNT25	SipL	134±17		134±14
		-				110±23
	CotL + CotB	pUT18C		pUT18
			CotL	CotB	CotL	CotB	-
		CotL		127±3		139±14
	pKT25	CotB	120±8		127±7
		CotL		151±7		947±428
	pKNT25	CotB	129±3		134±19
		-				93±45
	CotL + CotA	pUT18C		pUT18
			CotL	CotA	CotL	CotA	-
		CotL		164±48		114±34
	pKT25	CotA	88±31		113±61
		CotL		76±28		140±10
	pKNT25	CotA	ND		ND
		-				124±6
	CotL + CotE	pUT18C		pUT18
			CotL	CotE	CotL	CotE	-
		CotL		112±86		69±32
	pKT25	CotE	125±14		53±5
		CotL		161±151		324±153
	pKNT25	CotE	143±16		128±27
		-				106±42
	CotL + CdeC	pUT18C		pUT18
			CotL	CdeC	CotL	CdeC	-
		CotL		128±17		134±13
	pKT25	CdeC	71±18		135±21
		CotL		172±28		144±17
	pKNT25	CdeC	144±25		138±18
		-				110±23
	CotL + CotL	pUT18C	pUT18
			CotL	CotL	-
	pKT25	CotL	101±52	107+44
		CotL	98±45	211±11
	pKNT25	-			126±3

Values shown are the average ß-galactosidase activities (in Miller unit) of the indicated pair-wise fusions. Each value corresponds to the mean and the standard deviation of 4 or 5 independent experiments from double transformants. Positive interactions are indicated in orange.

Table 2 . Comparative analysis of the spore coat/exosporium proteome from the wild- type and the sigB mutant.

 2 Relevant proteins are indicated with an asterisk.

	Protein (Gene)	MW	Function	Fold-	-LOG(p-	Know/predicted
		[kDa]		change	value)	σ factor needed
				(WT/CD106		for expression
				5)
	Down in the sigB
	mutant
	CD0174	68.1	Carbon monoxide dehydrogenase	12.4	6.4	σ B
			cooS
	CD0175	17.8	oxidoreductase, Fe-S subunit	6.6	3.8	σ B
	CD0176*	45.7	Oxidoreductase, NAD/FAD	7.1	2.3	σ B
			binding subunit
	CD1125*	32.7	Nitroreductase-family protein	4.6	3.8	σ B
	CD1474*	20.0	Reverse Rubrerythrin	7.6	4.6	σ B
	CD1524*	20.0	Reverse Rubrerythrin	2.6	2.3	σ B
	CD1623*	94.3	Oxidoreductase	4.3	6.0	σ B
	CD2373	53.0	Carbon starvation protein	3.2	3.4	σ B
	CD2820	17.7	Hypothetical protein	4.6	2.1
	CD2996	27.4	Hypothetical protein	3.4	2.8
	CD3614	24.0	Hypothetical protein	4.1	1.8
	CD1176	6.5	Uncharacterized protein	3.9	2.0
	CD1404	66.6	Oligopeptide transporter	3.5	3.8
	CD1479	78.2	Ferrous iron transport protein B	3.8	4.3
	Up in the sigB
	mutant
	CD1396	29.7	Amino acid amidase	0.1	3.7
	CD1543	8.1	FMN-dependent NADH-	0.1	2.5	σ G
			azoreductase
	CD2575	24.2	Ribulose-phosphate 3-epimerase	0.2	2.3
	CD2841	60.2	Putative amidohydrolase	0.3	2.4	σ B
	CD2845*	22.4	Rubrerythrin	0.3	2.5	σ G
	CD3490	88.5	Stage II sporulation protein E	0.4	3.3	σ A
			SpoIIE

Table S2 : Oligonucleotides used in this study.

 S2 Restriction sites and tales used for Gibson assembly clonning are underlined.

			Results Results
	CF118	CCGCTCGAGTTCATTATCTTCTACCTCCTCTAT CC	Complementation CD2399 comp vector
	IMV918	GTAGAAATACGGTGTTTTTTAGCACAACTAACACTATAA	3' XhoI ACE CD1474 3' PCR2 rev
	CF211	CGCGGATCCTATGAAAAAATTAGATACAGA TTC	BACTH 5' cotL BamHI comp vector
	CF212 IMV919	CGAATTCCTATTCATGAGGGTTAGACTT TCATGAGATTATCAAAAGGAAGGAGGGGGCTTTATTGA	BACTH 3' cotL EcoRI ACE CD1524 5' PCR1 rev
		AA	pKT25, pUT18C comp vector
	CF213 IMV920	CGAATTCGATTCATGAGGGTTAGACTTT GTAGAAATACGGTGTTTTTTGTTCATGGCAACAGTAACA	BACTH 3' cotL EcoRI ACE CD1524 3' PCR2 rev
		TC	pNT25, pUT18 comp vector
	CF217 IMV949	CGCGGATCCTATGATAGATAATCAAAAATATGT AGGAATTCCTCCAAACAAGCTCTT	BACTH 5' cotB BamHI 5' Pσ A
	Primer CF218	Sequence (5' to 3') a CGAATTCTTACATGTTTTTATAACTCTCCAATATT	Description BACTH 3' cotB EcoRI
	CF14 CF219	CGCGGATCCACCCTTTCCAATTGTTTATAGTG CGAATTCGACATGTTTTTATAACTCTCCAATATT	ACE 5' CD2399 PCR1 pKT25, pUT18C BamHI BACTH 3' cotB EcoRI
	CF15	GGAATTCCTCCTTGTTGTAAATAGATAGGTAA	ACE 3' CD2399 PCR1 pNT25, pUT18
	CF220	CGCGGATCCTATGGAATTAATTAAAGATGTAAT	EcoRI BACTH 5' sipL BamHI
	CF16 CF221	GGAATTCCTAGAGTCTGAAATGTACAACAAAG CGAATTCTTAATCTACTAATACGACTTTTTTTTC	ACE 5' CD2399 PCR2 BACTH 3' sipL EcoRI
			EcoRI pKT25, pUT18C
	CF17 CF222	CCGCTCGAGTTACTATTTGATTGATACCTGCTT CGAATTCGAATCTACTAATACGACTTTTTTTTCT	ACE 3' CD2399 PCR2 BACTH 3' sipL EcoRI
			XhoI pNT25, pUT18
	CF19 CF223	GGAATTCGCTCATAGATTAAGTACCATATC CGCGGATCCAATGCAAGATTATAAAAAAAATAAA	5' CD1474 promoter EcoRI BACTH 5' cdeC BamHI
	CF20 CF224	CCGCTCGAGAATTTT TTATTTATTAAGTTAGTC CGAATTCCTATCTGTGGCAACTTGGCT	3' CD1474 promoter XhoI BACTH 3' cdeC EcoRI
	CF21	GGAATTCGTT TAG ACA AAG ATT TAG ATG TTA C	5' CD1623 promoter EcoRI pKT25, pUT18C
	CF22 CF225	CCGCTCGAG TAA AAT AGT TAT TTA GTC TCT T CGAATTCGATCTGTGGCAACTTGGCTT	3' CD1623 promoter XhoI BACTH 3' cdeC EcoRI
	CF23	GGAATTCCAGAGGAGTCTTCATTAGAAA G	5' CD1524 promoter EcoRI pNT25, pUT18
	CF24 CF226	CCGCTCGAGAATTTTTAT TTTTAAGTTTAATA CGCGGATCCGGTGATTTACATGCCAAA	3' CD1524 promoter XhoI BACTH 5' cotE BamHI
	CF25 CF227	GGAATTCGTATTG GATGGAAGTGAGAAGGAT CGAATTCTTAGAATTGCCCATAAATACCTT C	5' CD0176 promoter EcoRI BACTH 3' cotE EcoRI
	CF26	CCGCTCGAGTATTTTTATATTAAAAAAGTAGAT TA	3' CD0176 promoter XhoI pKT25, pUT18C
	CF30 CF228	TAT AAA TAT CAA AAC CAA CAT TA CGAATTCGAGAATTGCCCATAAATACCTTCA	CD1474 sequencing BACTH 3' cotE EcoRI
	CF31	TTA TTA GTG GTA AGT ATG CTT ATA	CD1623 sequencing pNT25, pUT18
	CF39 CF229	TTC TTC AAT TCC AGA TGC CT CGCGGATCCCGTGGAAAATAATAAATGTA	CD1524 sequencing BACTH 5' cotA BamHI
	CF95 CF230	CGTAATCATGGTCATATGGATA CGAATTCTTATGCAATATAATCTATAGAATCTAC	pFT58-adjacent EcoRI BACTH 3' cotA EcoRI
	CF96	GGATCCGCAGCTGCTGATAAAGA	pFT58-adjacent BamHI pKT25, pUT18C
	CF99 CF231	TATCCATATGACCATGATTACGGCTCATAGATTAAGTAC CGAATTCGATGCAATATAATCTATAGAATCTACA	5' CD1474 promoter BACTH 3' cotA EcoRI
		CATATC	pNT25, pUT18
	CF100 CF236	TCTTTATCAGCAGCTGCGGATCCTTTTCCAAAATATCTA GGAATTCTATGAAAAAATTAGATACAGA	3' CD1474 petDuet 5' cotL EcoRI
	CF130	TTTAATAATC ATAAGAATGCGGCCGCCTATTCATGAGGGTTAGACTTTT	petDuet 3' cotL NotI
	CF101	TATCCATATGACCATGATTACGGTTTAGACAAAGATTTA T	5' CD1623 promoter
	CF184	GATG TTAC GGAATTCCATATGAAAAAATTAGATACAGAGTCA	petDuet 5' cotL NdeI
	CF102 CF237	TCTTTATCAGCAGCTGCGGATCCTAGTATATCAGCCTTC CCGCTC GAGCTATTCATGAGGGTTAGACTTTTT	3' CD1623 petDuet 3' cotL XhoI
	CF238	ATAAT G GAATTCTATGATAGATAATCAAAAATATGT	petDuet 5' cotB EcoRI
	CF103 CF239	TATCCATATGACCATGATTACGGTATTGGATGGAAGTGA ATAAGAATGCGGCCGCTTACATGTTTTTATAACTCTCCA	5' CD0176 promoter petDuet 3' cotB NotI
		GAAGGAT ATATT
	CF104 CF244	TATTTTTATATTAAAAAAGTAGATTA GGAATTCGGTGATTTACATGCCAAA	3' CD0176 promoter petDuet 5' cotE EcoRI
	CF105 CF245	TAATCTACTTTTTTAATATAAAAATACAGAAGGAGGAA ATAAGAATGCGGCCGCTTAGAATTGCCCATAAATACCTT	5' CD0176 petDuet 3' cotE NotI
		AATGATGAATTA C
	CF106 CF248	TCTTTATCAGCAGCTGCGGATCCTACACTATAACAAAAT GGAATTCTATGGAATTAATTAAAGATGTAATTA	3' CD0176 petDuet 5' sipL EcoRI
	CF249	TCTCC ATAAGAATGCGGCCGCTTAATCTACTAATACGACTTTTT	petDuet 3' sipL NotI
	CF107	TATCCATATGACCATGATTACGTATACACGATAAAATAT TT	5' CD1157 promoter
	CF254	TTTCTAT GGAATTCCATATGTATAGGAGGAATAATATGAAT	petDuet 5' spoIVA NdeI
	CF108 CF255	TCTTTATCAGCAGCTGCGGATCCTTTAACTAATTTTGCA CCGCTC GAG TTATAACAAAATAGTTATAATATTAGA	3' CD1157 petDuet 3' spoIVA XhoI
	IMV699	AATTCTGC CCGCTCGAGAAAAGGGAACTGTATTG	Complementation CD1623
	CF113	TCCCCCCGGGCAGAGGAGTCTTCATTAGAAAG	Complementation CD1524 5' XhoI
	IMV700	CGGGATCCTTAAACCTATGTATGTA	5' XmaI Complementation CD1623
	CF114	CCGCTCGAGTTATTTTCCAAAATGTCTGTTTAA	Complementation CD1524 3'BamHI
	IMV844	CTTTACTGGAGTAGTAGATGAA	3' XhoI Verification intron CD1157
	CF115 IMV845	GGAATTCGCTCATAGATTAAGTACCATATC GACCATCTAAGTATAGTTTAGCA	Complementation CD1474 Verification intron CD1157
	IMV914	TCATGAGATTATCAAAAGGGGACAATTTATCTTGATTTA	5' EcoRI ACE CD1623 5' PCR1 rev
	CF116	ATAAGAATGCGGCCGCTTATTTTCCAAAATATCTATTTA CTAG	Complementation CD1474 comp vector
	IMV915	ATAATC GTAGAAATACGGTGTTTTTTCACTTGAGTAAAGTCAGGA	3' NotI ACE CD1623 3' PCR2 rev
	CF117	GGAATTCTTTACAAAAAAGTCTACCTTCATC TT	Complementation CD2399 comp vector
	IMV917	5' EcoRI TCATGAGATTATCAAAAGGGATAGACTTCTAAATATTTC ACE CD1474 5' PCR1 rev
		138 139

a

  . However, the in vivo results obtained in this work indicate that these proteins confer tolerance to O 2 , and this protection most likely arises from their ability to scavenge O 2 . A C. difficile double mutant inactivated for the two revRbrs and a mutant inactivated for the FDP CD1157 showed decreased tolerance to low O 2 tension. Accordingly, our complementary in vitro results showed that the revRbrs, CD1474 and CD1524, or the FDP CD1157 from C. difficile, in association with a NROR and a Rd from E. coli, function as NADH oxidase, reducing O 2 to water. Interestingly, the genes encoding the two revRbrs, Rbr3B and Rbr3A, and the two FDPs, FprA1 and FprA2, from C. acetobutylicum are upregulated after O 2

Table 3 . Reductase activities of the two revRbrs CD1474 and CD1524, and the two FDPs CD1157 and CD1623 to the substrates O 2 , H 2 O 2 and NO.

 3 acetobutylicum, the C. difficile revRbrs can use both O 2 and H 2 O 2 as electron acceptors, but H 2 O 2 is the preferred substrate. For both activities, the electron transfer pathway did not absolutely require the Rd from E. coli, but its presence increased the NADH peroxidase and oxidase activities. The physiological electron donors of both revRbrs and the FDP CD1157 from C. difficile remain unknown. However, since the genome of the C. difficile 630 strain CD0176 deletion mutant. However, the inactivation of this gene did not result in a clear effect on the sensitivity of C. difficile to several stresses such as O 2 or H 2 O 2 . The enzymatic role of CD0176 is currently under study by our collaborators in Portugal. In addition, an interesting

	does not contain genes encoding Rds, the NAD(P)H-oxidoreductases (ORs) in this strain
	most likely directly reduce the [Fe(SCys) 4 ] site of the revRbrs or the FMN domain of the
	FDP. Possible candidates for ORs in C. difficile might be the CD0176, the NADH oxidase
	CD2540 and a NADH-dependent flavin OR CD2709. In the laboratory, we have constructed a

  KatA) or σ B (KatB and KatX). In this way, the response induced by oxidative stress would be either specific through PerR or mediated by a general stress response via σ B(Hecker et al. 2007). In C. difficile, PerR and σ B are present like in B. subtilis. However no induction of σ B activity has been observed to date in the presence of ROS and the H 2 O 2 protection mediated by σ B is probably unspecific. Concerning the role of PerR and its targets in C. difficile, nothing has been yet studied, as the effect of H 2 O 2 on gene expression. Further work will be needed to determine the targets of PerR and the connection that both PerR and σ B might have to protect this bacterium from oxidative stress.

	Discussion
	#1
	O 2 by either PerR (
	NO
	O 2 / NO
	…....
	#2
	O 2
	NO
	O 2 / NO
	Figure 60: Amino acid sequence alignment of FDPs. The C.difficile FDPs CD1157 and CD1623
	were aligned against the FDPs from Thermotoga maritima, Entamoeba histolytica, Giardia intestinalis,
	Methanothermobacter marburgensis, Escherichia coli, Moorella thermoacetica and Desulfovibrio
	gigas. Green arrows indicate the residues in positions 1 and 2. The selectivity for O 2 or NO is also
	indicated.
	156

Table S2 : Oligonucleotides Name Sequence

 S2

			Annexes Annexes
	NK47	TAGACATGTAAAAACCTCCTATTTCCTTTT	DCO rsbV 3'PCR1 pUT18/pKNT25
	NK48 NK120	AAAAGGAAATAGGAGGTTTTTACATGTCTAAAGAAAGA GATAGAATTCCATTAAATATCAACTCCTAAATAT	DCO rsbV 3'PCR2 rev BACTH 3' EcoRI RsbW
		TTTTTACTATAACAGGTCTAG	comp NK47 pUT18C/pKT25
	NK49 NK121	ATCGTAGAAATACGGTGTTTTTTCTTGTTCCTTTTTTCAA GATAGGATCCGTCTATGAATATTGATTCAAATTTAG	DCO rsbV 5'PCR2 BACTH 5' BamHI RsbV
		GGAG	pUT18/pUT18C	and
	NK129	GATCGGATCCCTACCTATCTTGCATACCAACAGAAACTT	3' pKNT25/pKT25 rsbZ ∆phosphatase
	NK122	G GATAGAATTCCCTCCCTCCACTTTAAATATTT	domain BamHI BACTH 3' EcoRI RsbV
	NK130	GCAATAAAAGAAGTTGTAAAAAAGGGGAAATAGGAGG	DCO CD0007-08 PCR2 pUT18/pKNT25
	NK123	TTTTTACATGTCTA GATAGAATTCGTTTATCCCTCCACTTTAA	5'rev comp NK39 BACTH 3' EcoRI RsbV
	NK108	TGGTCATGAGATTATCAAAAGGGATAAACTAAGAGGT	DCO ptet CD2684 5' PCR1 pUT18C/pKT25
	NK183	TCAATAAG GATAGGATCCATT GATTATGGATTCAAACGATA	BACTH 5' BamHI CD2684
	NK109	ATCGTAGAAATACGGTGTTTTTTTAGTCGCGCAAATACT CTATC	pUT18/pUT18C DCO ptet CD2684 3' PCR2 and pKNT25/pKT25
	NK228 NK184	CGTTAACAGATCTGAGCTCCTTGATTATGGATTCAAACG ATAAAA GATAGAATTCAATTATATTGAATCAATATATATC	BACTH 3' EcoRI CD2684 DCO ptet CD2684 5' PCR2 pUT18C/pKT25
	NK229 NK185	CTAGCTTGATGCAGAATTCGTCTACCTATCCTTTAACAT AAC GATAGAATTCCCTATTGAATCAATATATATCTTTAA	BACTH 3' EcoRI RsbV DCO ptet CD2684 3'PCR1 pUT18/pKNT25
	NK230 NK124	TTTTATCGTTTGAATCCATAATCAAGGAGCTCAGATCTG CAATTTCACACAGGAAACAGC	DCO ptet CD2684, ptet BACTH 5' pUT18/pUT18C
	NK125	TTAACG GAGCAGATTGTACTGAGAG	element 3' BACTH 3' pUT18/pUT18C
	NK231 NK126	GTTATGTTAAAGGATAGGTAGACGAATTCTGCATCAAG CGGATAACAATTTCACACAGG	DCO ptet CD2684, ptet BACTH 5' pKT25/pKNT25
	NK127	CTAG GAGAGTGCACCATATTACTTAG	element 5' BACTH 3' pKNT25
	NK148 NK128	GATCAGGCCTGGAAATAGGAGGTTTTTACATGTCTA GTTGTAAAACGACGGCCG	5' StuI rsbV BACTH 3' pKT25
	NK149 IMV388	GATCCTCGAGCTTTATAGTCTCACAAGCCAAAG GAGTCCCAAGAAGAATACAGGA	3' XhoI rsbV qRT-PCR sigB
	NK155 IMV389	GATCAGGCCTATGGATAAAGATTGTGAAATGA AAGCCTCCATAGCCTCTAAAAC	StuI 5' SNAP Cd (ORF) qRT-PCR sigB
	NK156 IMV774	GATCCTCGAGTTATTCCTTACCCAAGTCCT CCAAAAGAAGTTCTACATGAAACA	XhoI 3' SNAP Cd (ORF) qRT-PCR CD0007
	NK181 IMV775	GATCGGATCCTTATTCCTTACCCAAGTCCT AACATCTCCCTCATCATCAAA	BamHI 3' SNAP Cd (ORF) qRT-PCR CD0007
	NK210 IMV776	GATCGGATCCCTTTATAGTCTCACAAGCCAAAG TGGAATGTTTGTCTTGATGGA	3' RsbV BamHI qRT-PCR rbsV (CD0009)
	NK211 IMV777	GATCAGGCCTAAGAAAGATTTTTACTATAACAGG AGCTCCTAAGCCTGTTGAGTCT	5' RsbW StuI qRT-PCR rbsV (CD0009)
	NK13 NK212 IMV778	GATCAGGCCTGTTAAAATACTAATAAATAGTT GATCGGATCCGCTCTTTTGTATCCATATTTAGG TCCAAAAGAGAGTGGTCTTGG	5' CD2685 StuI 3' RsbW BamHI qRT-PCR rbsW (CD0010)
	NK14 NK194 IMV779	GGGATCCTAAGTCCTTTTTTATCG GTTCCAGATTACGCTTAGGGATCCTATAAGTTTTAAT AGTCATTTTTATTTTTGTCCCTTG	3' CD2685 BamHI RsbZ ∆pho HA C-ter inverse qRT-PCR rbsW (CD0010)
	NK15 OS547	AGGCCTAAAGATAATTATGATGATG GGCAAATTGGAGCAGTGTTA	5' CD2684 StuI PCR 1 qRT-PCR CD0008
	NK16 NK195 OS548	GGGATCCCGAATACAATTATACATAAT ATCGTATGGGTACCTATCTTGCATACCAACAGA TCTTATATGCCTTTTCAACAACCTT	3' CD2684 BamHI RsbZ ∆pho HA C-ter inverse qRT-PCR CD0008
	NK17 NK30 NK18 NK192 NK31 NK19 IMV846 NK20 NK21 NK22 NK23 NK24 NK104 NK39 NK40 NK193 IMV847 NK188 IMV668 IMV669 NK189 IMV670 IMV671 NK112 IMV575 IMV576 IMV747 NK113 IMV748 IMV790 NK114 IMV791	AGGCCTTTTGTATAATGTAGTTAGATA AGAGCAGAAGGTGAACGAGA GGGATCCAGCTTCCTACTAAC GTTCCAGATTACGCTTAAACTTTGGCTTGTGAGAC TCAAGAGTGATGCAACAGATGA AGGCCTGTAATAAGTAAAAGCTTTG TTGCACAGCATGTGGATTTT GGGATCCTCTAAATTTGAATCAATA CTCGAGTTATTTGAGAGATATTGAAAC TCATACTCAATTTCTCCCCCTA TAGGGGGAGAAATTGAGTATGAGGTATAAACCGATTAG AAAACTTT GGGATCCAACTTGAGTATGAGCTCTA TGGTCATGAGATTATCAAAAGGACTGAAATCCAAGCAC AAGC CCACCTTTTTTACAACTTCTTTTATTGC GCAATAAAAGAAGTTGTAAAAAAGGTGGGCTTTGATGA TGAGGGAGATGTTAAC ATCGTATGGGTATCCCTCCACTTTAAATATTTTAT CCATCACAATCTGTCCACCTT GTTCCAGATTACGCTTAAAAAAATATAAAAAAAGTATT GACC TGCCCTGTATGTGGAGCTAA CCAGCTGCTCCATTTCCTAC ACATGAAGGAGATGCTGCAC ATCGTATGGGTATAAATTTTTTTCATATTCTTTTTTCAGC SigB HA C-ter inverse PCR PCR 2 qRT-PCR CD2685 (rsbZ) 5' CD0007 StuI RsbV HA C-ter inverse PCR qRT-PCR CD2685 (rsbZ) 3' CD0007 BamHI 1 qRT-PCR CD2684 5' CD0008 StuI 3' CD0008 BamHI 5'CD2685 DCO XhoI PCR1 3' CD2685 DCO PCR1 5' CD2685 DCO PCR2-rev-comp NK22 3' CD2685 DCO BamHI PCR2 DCO CD0007 5'PCR1 DCO CD0007 3'PCR1 DCO CD0007 3'PCR2 rev pKT25/pUT18C comp NK39 RsbV HA C-ter inverse PCR qRT-PCR CD2684 2 qRT-PCR CD1623 SigB HA C-ter inverse PCR qRT-PCR CD1623 1 qRT-PCR CD1474/CD1524 GCTGAACATGCTGCTAAATTT qRT-PCR CD1474/CD1524 2 GATAGGATCCAATGAAAAATGTAGCTAATGC CCAAGGCAAGATAGCAGGAG qRT PCR CD1157-norV BACTH BamHI sigB 5' pKNT25/pKT25 CTGCACCAAATGCCATACAC qRT PCR CD1157-norV and GGGGTTAAACCAAATGCTGA qRT-PCR CD0176 PUT18/pUT18C GATAGAATTCCCTAAATTTTTTTCATATTCTTTT CCTTGCTTTACTGCCAGAGG qRT-PCR CD0176 BACTH EcoRI sigB 3' GCTACAAAAACTGGGCAAAA qRT-PCR CD1125 pKNT25/pUT18 GATAGAATTCTTTTATAAATTTTTTTCATATTC BACTH EcoRI sigB 3' ACATTTCCCCATTCGTTCAT qRT-PCR CD1125
	NK105 NK115	ATCGTAGAAATACGGTGTTTTTTTCAACATCATATCTAT GATAGGATCCATTGAGTATGACTAATGGCAG	DCO CD0007 5'PCR2 BACTH 5' BamHI CD2685
		CAATAGC	pKNT25/pKT25	and
	NK106	TGGTCATGAGATTATCAAAAGGAAGAAGATTTTGTAAG	DCO CD0008 5'PCR1 pUT18C/pUT18
	NK116	GCACTTG GATAGAATTCCCCCTATCCTTTAACATAACTATT	BACTH 3' EcoRI CD2685
	NK43	AATTTGCCACCCCCATGCATTCA	DCO CD0008 3'PCR1 pKNT25/pUT18
	NK44 NK117	TGAATGCATGGGGGTGGCAAATTGGAAATAGGAGGTTT GATAGAATTCATCTACCTATCCTTTAACATAAC	DCO CD0008 3'PCR2 rev BACTH 3' EcoRI CD2685
		TTACATGTCTA	comp NK43 pKT25/pUT18
	NK107 NK118	ATCGTAGAAATACGGTGTTTTTTTTCGCATCACTTATCTT GATAGGATCCTTTGGCTTGTGAGACTATAAA	DCO CD0008 5'PCR2 BACTH 5' BamHI RsbW
		TTTGC	pUT18/pUT18C	and
	NK110	TGGTCATGAGATTATCAAAAGGGACTAAGAGAGAAGAG	DCO rsbV 5'PCR1 pKNT25/pKT25
	NK119	TTCAA GATAGAATTCCCAATATCAACTCCTAAATATTTAG	BACTH 3' EcoRI RsbW
		226 227

Supplementary tables of the Article: CotL, a new morphogenetic spore coat protein of Clostridium difficile Table S1. Bacterial strains and plasmids used in this study.

						Annexes
	CDIP1067	630Δerm sigE::erm pDIA6663			This work
	CDIP1068	630Δerm sigK::erm pDIA6663			This work
	Plasmids	Relevant properties				Origin
	pET20	E. coli expressing vector (Amp R )
	pDIA6636	pET20-CD1065-His 6				This work
	pMTL007	group II intron, ErmBtdRAM2 and ltrA of pMTL20lacZ-	Heap et al., 2007
		TTErm ;BtdRAM2, Cm R /Tm R
	pDIA6563	pMTL007-CE5-CD1065-123s (Cm R /Tm R )		This work
	pMTL84121	Clostridium modular plasmid containing catP (Cm R /Tm R )	Heap et al., 2009
	pFT47	(Cm R /Tm R ) pMTL84121-SNAP Cd	(for	transcriptional	fusions)	Pereira et al., 2013
	Strains E. coli pFT58	Genotypes (Cm R /Tm R ) pMTL84121-linker-SNAP Cd (for translational fusions)	Origin Pereira et al., 2013
	10-beta pDIA6626 pDIA6561 pDIA6613 HB101 (RP4) PDIA6617 pDIA6631 pDIA6633	F -mcrA D(mrr-hsdRMS-mcrBC) f80lacZDM15 DlacX74 pMTL84121-P σE/σK(CD1065) -CD1065 pFT47 P σE/σK(CD1065) -SNAP Cd deoR, recA1 araD139 D(ara-leu)7697 galK rpsL(StrR) pFT47 P σK(CD1065) -SNAP Cd endA1 nupG supE44 aa14 galK2 lacY1 D(gpt-proA) 62 rpsL20 (Str R ) xyl-5 mtl-1 recA13 D(mcrC-mrr) hsdS B (r B -) RP4 (Tra + pFT58 P σE/σK(CD1065) -CD1065-SNAP Cd pFT58 P spoIVA -spoIVA-SNAP Cd -m B IncP Ap R Km R Tc R ) pFT58 P cotB -cotB-SNAP Cd	This work NEB This work This work This work Laboratory stock This work This work
	BL21 (DE3) pDIA6634	fhuA2 (lon) ompT gal (λ DE3) (dcm) ΔhsdS λ DE3= λ pFT58 P cotE -cotE-SNAP Cd	NEB This work
	pDIA6632	sBamHIo ΔEcoRI-B int::(lacI::PlacUV5::T7 gen1) i21 pFT58 P cdeC -cdeC-SNAP Cd	This work
	pDIA6661	Δnin5 / E.coli strain for T7 expression, protease deficient pFT58 P cotD -cotD-SNAP Cd	This work
	pDIA6663	B strain pFT58 P σK(CD1065) -CD1065-SNAP Cd		This work
	ECO832 pDIA6664	BL21 (DE3) pET20-CD1065 pFT47 P σE(CD1065)-SNAP Cd			This work This work
	C. difficile
	630∆erm					Laboratory stock
	AHCD532	630Δerm sigE::erm				Pereira et al., 2013
	AHCD535	630Δerm sigK::erm				Pereira et al., 2013
	CDIP734	630Δerm pFT47				This work
	CDIP782	630∆erm CD1065::erm				This work
	CDIP753	630Δerm pFT58				This work
	CDIP851	630∆erm pDIA6617				This work
	CDIP871	630∆erm pMTL84121				This work
	CDIP873	630∆erm CD1065::erm pMTL84121		This work
	CDIP881	630∆erm pDIA6561				This work
	CDIP882	630Δerm sigE::erm pDIA6561			This work
	CDIP883	630Δerm sigK::erm pDIA6561			This work
	CDIP948	630Δerm sigE::erm pDIA6617			This work
	CDIP950	630Δerm sigK::erm pDIA6617			This work
	CDIP952	630∆erm CD1065::erm pDIA6626		This work
	CDIP986	630∆erm pDIA6613				This work
	CDIP987	630Δerm sigE::erm pDIA6613			This work
	CDIP989	630Δerm sigK::erm pDIA6613			This work
	CDIP1070	630∆erm pDIA6664				This work
	CDIP1072	630Δerm sigE::erm pDIA6664			This work
	CDIP1071	630Δerm sigK::erm pDIA6664			This work
	CDIP1035	630∆erm pDIA6634				This work
	CDIP1039	630∆erm CD1065::erm pDIA6634		This work
	CDIP1036	630∆erm pDIA6632				This work
	CDIP1040	630∆erm CD1065::erm pDIA6632		This work
	CDIP1037	630∆erm pDIA6633				This work
	CDIP1041	630∆erm CD1065::erm pDIA6633		This work
	CDIP1038	630∆erm pDIA6631				This work
	CDIP1042	630∆erm CD1065::erm pDIA6631		This work
	CDIP1057	630∆erm pDIA6661				This work
	CDIP1058	630∆erm CD1065::erm pDIA6661		This work
	CDIP1066	630∆erm pDIA6663				This work
						235

Table S4 : Effect of CD1065 inactivation on the expression of sporulation- specific genes.

 S4

Acknowledgments Acknowledgements

We thank Maryse Moya-Nilges for performing transmission electron microscopy, Jost Enniga and Giulia Manina for access to the fluorescence microscope, Adriano Henriques for providing the plasmids carrying the SNAP Cd reporter gene and the antibody raised against CdeM, Aimee Shen for helpful discussion and providing the antibodies, Nigel Minton for the ClosTron system and Julian Garneau for the bioinformatics analysis. Research in this study for the CAF fellowship was funded by the Institut Pasteur and ITN Marie Curie, Clospore (H2020-MSCA-ITN-2014 642068). T. maritima E. histolytica G. intestinalis E. coli D. gigas C. difficile CD1157 C. difficile CD1623 M. marbugensis M. termoacetica T. maritima E. histolytica G. intestinalis E. coli D. gigas C. difficile CD1157 C. difficile CD1623 M. marbugensis M. termoacetica

Acknowledgments

We are thankful to Johann Peltier and Gouzel Karimova for helpful discussions, to Jean-Marc Ghigo for his help with oxygen assays, to Julian Garneau for his help with the bioinformatic analysis of sigB and rsbZ operon and to Claire Morvan for a critical reading of

ECO1033 BL21 (DE3) + pDIA6840 This work ECO1035 BL21 (DE3) + pDIA6841 This work ECO1036 BL21 (DE3) + pDIA6842 This work ECO1037 BL21 (DE3) + pDIA6843 This work ECO1038 BL21 (DE3) + pDIA6844 This work ECO1039 BL21 (DE3) + pDIA6845 This work ECO1041 BL21 (DE3) + pDIA6846 This work ECO1042 BL21 (DE3) + pDIA6847 This work

C. difficile CDIP546 630∆erm sigB::erm Kint et al, 2017 CDIP588 630∆erm CD1157::erm This work CDIP658 630∆erm + pDIA6458 This work CDIP659 630∆erm + pDIA6459 This work CDIP661 630∆erm + pDIA6457 This work CDIP666 630Δerm + pDIA6460 This work CDIP668 630∆erm sigB::erm + pDIA6457 This work CDIP669 630∆erm sigB::erm + pDIA6458 This work CDIP670 630∆erm sigB::erm + pDIA6459 This work CDIP671 630∆erm sigB::erm + pDIA6460 This work CDIP691 630Δerm ΔCD1524 This work CDIP697 630Δerm ΔCD1474 This work CDIP714 630∆erm ∆CD1524 ∆CD1474 This work CDIP717 630 Δerm ΔCD2399 This work CDIP734 630∆erm pFT47 This work CDIP740 630∆erm sigB::erm + pDIA6497 This work CDIP742 630∆erm + pDIA6503 This work CDIP743 630∆erm sigB::erm + pDIA6503 This work CDIP744 630∆erm + pDIA6499 This work CDIP745 630∆erm sigB::erm + pDIA6499 This work CDIP774 630∆erm sigB::erm + pDIA6517 This work CDIP750 630∆erm + pDIA6497 This work CDIP751 630∆erm + pDIA6498 This work CDIP752 630∆erm sigB::erm + pDIA6498 This work CDIP753 630∆erm pFT58 This work CDIP767 630∆erm + pDIA6517 This work CDIP798 630∆erm ∆CD2399 + pDIA6534 (complementation) This work CDIP799 630∆erm ∆CD1524 + pDIA6537 This work CDIP800 630∆erm ∆CD1524 ∆CD1474 + pDIA6537 This work CDIP801 630∆erm ∆CD1474 + pDIA6538 (complementation) This work CDIP802 630∆erm ∆CD1524 ∆CD1474 + pDIA6538 This work CDIP843 630∆erm + pDIA6589 This work CDIP845 630∆erm sigB::erm + pDIA6589 This work CDIP1045 630∆erm + pDIA6669 This work CDIP1046 630∆erm + pDIA6670 This work CDIP1047 630∆erm sigB::erm + pDIA6669 This work CDIP1048 630∆erm sigB::erm + pDIA6670 This work CDIP1136 630∆erm cdeM::erm + pDIA6617 This work CDIP1137 630∆erm cdeM::erm + pDIA6663 This work CDIP1369 630Δerm ΔCD1623 This work pDIA6374 pMTL007-CD1157 Cm R -Tm R This work pDIA6427 pMTL-SC7315 ∆CD2399 This work pDIA6429 pMTL-SC7315-∆CD1524 Cm R -Tm R This work pDIA6457 pFT47 P CD1474 -SNAP Cd This work pDIA6458 pFT47 P CD1623 -SNAP Cd This work pDIA6459 pFT47 P CD1524 -SNAP Cd This work pDIA6460 pFT47 P CD0176 -SNAP Cd This work pDIA6474 pMTL-SC7315-∆CD1474 Cm R -Tm R This work pDIA6497 pFT58 P CD1157 -SNAP Cd This work pDIA6498 pFT58 P CD1474 -SNAP Cd This work pDIA6499 pFT58 P CD1623 -SNAP Cd This work pDIA6503 pFT58 P CD0176 -SNAP Cd This work pDIA6517 pFT47 P CD1157 -SNAP Cd This work pDIA6534 pMTL84121-CD2399 Cm R -Tm R (complementation) This work pDIA6537 pMTL84121-CD1524 Cm R -Tm R (complementation) This work pDIA6538 pMTL84121-CD1474 Cm R -Tm R (complementation) This work pDIA6589 pFT47 P sigB -SNAP Cd This work PDIA6617 pFT58 P σE/σK(cotL) -cotL-SNAP Cd This work pDIA6635 pET20-CD1474 This work pDIA6663 pFT58 P σK(cotL) -cotL-SNAP Cd This work PDIA6669 P(σ B CD1157)-SNAP Cd This work pDIA6670 P(σ A CD1157)-SNAP Cd This work pDIA6671 pET20-CD1524 This work pDIA6700 pKT25-cotL This work pDIA6701 pKNT25-cotL This work pDIA6702 pUT18-cotL This work pDIA6703 pUT18C-cotL This work pDIA6704 pKT25-sipL This work Results 136 pDIA6705 pKNT25-sipL This work pDIA6706 pUT18-sipL This work pDIA6707 pUT18C-sipL This work pDIA6708 pKT25-cotB This work pDIA6709 pKNT25-cotB This work pDIA6710 pUT18-cotB This work pDIA6711 pUT18C-cotB This work pDIA6737 pKT25-cotE This work pDIA6738 pKNT25-cotE This work pDIA6739 pUT18-cotE This work pDIA6740 pUT18C-cotE This work pDIA6741 pKT25-cdeC This work pDIA6742 pKNT25-cdeC This work pDIA6743 pUT18-cdeC This work pDIA6744 pUT18C-cdeC This work pDIA6830 pUT18-cotA This work pDIA6831 pUT18C-cotA This work pDIA6832 pKT25-cotA This work pDIA6838 pETDuet His-cotE + cotL This work pDIA6839 pETDuet cotL This work pDIA6840 pETDuet His-cotL This work pDIA6841 pETDuet His-cotB + cotL This work pDIA6842 pETDuet His-sipL + cotL This work pDIA6843 pETDuet His-sipL This work pDIA6844 pETDuet His-cotL + spoIVA This work pDIA6845 pETDuet His-sipL + spoIVA This work pDIA6846 pETDuet His-cotE This work pDIA6847 pETDuet His-cotB This work pDIA6893 pMTL-SC7315-∆CD1623 Cm R -Tm R This work

CotL, a new morphogenetic spore coat protein of

Clostridium difficile

Carolina Alves Feliciano 1,2 , Thibaut Douché 3 , Quentin Giai Gianetto 3,4 , Mariette

Summary

The strict anaerobe Clostridium difficile is the most common cause of antibioticassociated diarrhoea. The oxygen-resistant C. difficile spores play a central role in the infectious cycle, contributing to transmission, infection and recurrence. The spore surface layers, the coat and exosporium, enable the spores to resist physical and chemical stress. However, little is known about the mechanisms of their assembly. In this study, we characterized a new spore protein, CotL, which is required for the assembly of the spore coat.

The cotL gene was expressed in the mother cell compartment under the dual control of the RNA polymerase sigma factors, σ E and σ K . CotL was localized in the spore coat, and the spores of the cotL mutant had a major morphologic defect at the level of the coat/exosporium layers. Therefore, the mutant spores contained a reduced amount of several coat/exosporium proteins and a defect in their localization in sporulating cells. Finally, cotL mutant spores were more sensitive to lysozyme and were impaired in germination, a phenotype likely to be associated with the structurally altered coat. Collectively, these results strongly suggest that CotL is a morphogenetic protein essential for the assembly of the spore coat in C. difficile.

Results

135

Summary

Clostridium difficile is the main cause of antibiotic-associated diarrhoea. Inside the gut, C. difficile must adapt to the stresses it copes with, by inducing protection, detoxification and repair systems that belong to the general stress response involving σ B . Following stresses, σ B activation requires a PP2C phosphatase to dephosphorylate the anti-anti-sigma factor

RsbV that allows its interaction with the anti-sigma factor RsbW and the release of σ B . In this work, we studied the signalling pathway responsible for the activation of σ B in C. difficile.

Contrary to other firmicutes, the expression of sigB in C. difficile is constitutive and not autoregulated. We confirmed the partner switching mechanism that involved RsbV, RsbW and σ B and we showed that CD2685, newly renamed RsbZ, and its phosphatase activity are required for RsbV dephosphorylation triggering σ B activation. While CD0007 and CD0008, whose genes belong to sigB operon, are not involved in σ B activity, depletion of the essential ironsulphur flavoprotein, CD2684, whose gene forms an operon with rsbZ, prevents σ B activation.

Finally, we observed that σ B is heterogeneously active in a subpopulation of C. difficile cells from the exponential phase, likely leading to a "bet-hedging" strategy allowing a better chance for the cells to survive adverse conditions.

Experimental procedures

Bacterial strains and growth conditions C. difficile strains and plasmids used in this study are presented in Table S1.

C. difficile strains were grown anaerobically (5% H 2 , 5% CO 2 , 90% N 2 ) in TY (Bacto tryptone 30 g.l -1 , yeast extract 20 g.l -1 , pH 7.4) or in Brain Heart Infusion (BHI; Difco). For solid media, agar was added to a final concentration of 17 g.l -1 . When necessary, cefoxitin (Cfx, 25 µg.ml -1 ), thiamphenicol (Tm, 15 µg.ml -1 ) or erythromycin (Erm, 2.5 µg.ml -1 ) were added to C. difficile cultures. E. coli strains were grown in LB broth. When indicated, ampicillin (100 µg.ml -1 ), chloramphenicol (15 µg.ml -1 ) or kanamycin (50 µg.ml -1 ) was added to the culture medium. The non-antibiotic analogue aTc was used for the induction of expression of the sigB, rsbV, rsbW or rsbZ (CD2685) genes expressed under the control of the P tet promoter that were cloned into pDIA6103 (Fagan and Fairweather 2011). Growth of C.

difficile strains in the presence of different concentration of aTc was followed using a GloMax® Explorer plate-reader (Promega). Overnight cultures were diluted at an OD 600nm of 0.05 in TY medium supplemented with 0, 10, 25, 50, 100 or 200 ng.ml -1 of aTc in 96 wells plates. The plates were sealed with a gas-impermeable adhesive film (DY992 R&D systems)

and OD 600nm was monitored every 30 min at 37°C.

Construction of C. difficile strains and plasmids

All routine plasmid constructions were carried out using standard procedure. All primers used in this study are listed in Table S2. ∆rsbV, ∆rsbZ, ∆CD0007, ∆CD0008 and ∆CD0007-0008 knock out mutant generated in this study were created using the codAmediated allele exchange method (Cartman et al. 2012, Peltier et al. 2015). Briefly, for all these genes, 1.5 kb fragments located up-and downstream of the gene to be deleted were PCR amplified from 630∆erm genomic DNA using primers NK46/NK47 and NK48/NK49 for ∆rsbV, NK21/NK22 and NK23/NK24 for ∆rsbZ, NK38/NK39 and NK40/NK41 for ∆CD0007, NK42/NK43 and NK44/NK45 for ∆CD0008 and NK38/39 and NK42/NK43 for ∆CD0007-0008 (Table S2). To construct CD2684 conditional mutant, two fragments of 1.5 kb corresponding to the end part of rsbZ and the entire CD2684 gene were amplified with NK108/NK229 and N228/NK109, respectively. A P tet and tetR gene fragment overlapping both the end of rsbZ in 5' extremity and the beginning of CD2684 in 3' extremity was amplified from pRPF185 plasmid with NK230 and NK231 in order to insert it right after the stop codon of rsbZ and before the initiation codon (TTG) of CD2684. Purified PCR fragments