Towards the understanding of the function and regulation of a ...

Towards the understanding of the

function and regulation of a membrane

protein complex involving SppA and YteJ

in Bacillus subtilis.

Thèse de doctorat de l'Université Paris-Saclay

préparée à l’Université Paris-Sud

École doctorale n°577 structure et dynamique des systèmes vivants (SDSV)

Spécialité de doctorat: Sciences de la vie et de la santé

Thèse présentée et soutenue à Jouy-en-Josas, le 1 Julliet 2019, par

Gabriela Fernandes Henriques

Composition du Jury:

Mme. Anne Galinier

Directrice de recherche, CNRS-IMM, Marseille Rapportrice

Mme. Nathalie Declerck

Directrice de recherche, INRA-CBS, Montpellier Rapportrice

Mme. Sylvie Nessler

Professeure, Université Paris-Sud XI, Orsay Président du Jury

Mme. Nathalie Campo

Chargée de recherche, CNRS-LMGM, Toulouse Examinatrice

M. Martin Picard

Directeur de recherche, CNRS- IBPC, Paris Examinateur

M. Jean Luc Pernodet

Directeur de Recherche, CNRS-I2BC, Orsay Examinateur

M. Matthieu Jules

Professeur, AgroParisTech, Micalis, Jouy-en-Josas Directeur de thèse

M. Olivier Delumeau

Ingénieur de recherche, Micalis, Jouy-en-Josas Co-Directeur de thèse

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Title : Towards the understanding of the function and regulation of a membrane complex

involving SppA and YteJ in Bacillus subtilis.

Keywords : Bacillus subtilis, Protein secretion, Membrane protein complex, Protease, Lantibiotic resistance,

Quality control

Abstract: Bacillus subtilis is a model organism for Gram-positive bacteria and a workhorse used for large-

scale production of industrial enzymes. Secretory and membrane proteins are mainly translocated through the

SecA-YEG (Sec) pathway. Although this highly conserved pathway has been extensively studied, there is a

lack of knowledge about the protein interactions and complexes involved in this mechanism in B. subtilis.

Therefore, to better understand these membrane protein complexes, we SPA-tagged a set of proteins involved

in the Sec pathway for tandem affinity purification. Through the combination of this technique with mass

spectrometry and Blue Native PAGE, we studied the protein interactions within the (holo)translocon as well

as with the accessory proteins. Our results showed that, under their SPA-tagged forms, SecDF and SpoIIIJ,

could not be retrieved together after immune-capture, as opposed to what was shown in E. coli. This result

indicated that SecDF and SpoIIIJ, unexpectedly, do not form a stable complex in B. subtilis. The other

experiments using SPA-tagged proteins (SecG, BdbB, and BdbC) did not provide evidence for the existence

of stable complexes involving proteins known to be part of the translocon complex, except for one protein, the

so-called signal peptide peptidase, SppA.Using the SPA-tag technique, we have identified a membrane protein

complex, involving SppA and another membrane protein, YteJ. While YteJ function is unknown, SppA is a

known membrane bound serine protease, described as responsible for recycling of the signal peptides of the

translocated proteins left in the membrane and also involved in the resistance to nisin, an antibacterial peptide

Of the lantibiotic family. Interestingly, sppA and yteJ genes are organized in operon, for which we confirmed

the regulation by two sigma factors, σA and σW, using transcriptional fusions. The fluorescent microscopy

experiments revealed a diffused membrane localization for both proteins, and therefore their co-localization

was difficult to prove by this technique. Using deletion mutant strains, we showed that both SppA and YteJ

are involved in the lantibiotic resistance: the sppA- strain was more sensitive while the yteJ- strain showed a

more sensitive phenotype than wild type, but less pronounced than the sppA- strain. The sppA-dependent

resistance was confirmed by the ectopic overexpression of SppA that restored the lantibiotic resistance.

However, the overexpression of SppA also induced the formation of elongated cells. Interestingly, this

phenotype can be suppressed by the simultaneous overexpression of YteJ, indicating an YteJ-dependent

negative control of SppA activity. Furthermore, using the elongated cell phenotype, we tested the

overexpression of SppA along with truncated versions of YteJ. The results pinpointed to the inhibitory role of

the C-terminal domain of YteJ. Finally, we performed in vitro biochemical studies with purified His-tagged

SppA and YteJ which showed that B subtilis SppA and YteJ form a membrane complex when expressed

together in E. coli. That the SppA protease activity of the purified complex His-SppA-YteJ was almost null,

and that, purified from the membrane, the full length SppA protease activity was lower than that of a soluble

truncated version of SppA. This full-length SppA activity was strongly reduced in the presence of YteJ,

supporting the hypothesis of an inhibition by YteJ. Furthermore, we showed that the ability or inability of

SppA to digest fully folded proteins probably depends on the structure of the proteins, suggesting a possible

role in protein quality control. In conclusion, our in vivo and in vitro studies showed that YteJ, via one of its

domain, acts as a negative regulator of the protease activity of SppA in this complex. Although we have

documented a role for this complex in the lantibiotic resistance, a possible role of SppA in protein quality

control might not be excluded

I

Titre : Caractérisation du complexe membranaire impliquant la signal peptide peptidase

SppA et YteJ chez Bacillus subtilis.

Mots clés : Sécretion protéique, complexe protéique membranaire, protéase, résistance aux

lantibiotiques, contrôle qualité

INTRODUCTION :

Les complexes membranaires impliqués dans la sécrétion des protéines chez B. subtilis ont été identifiés

mais ont été peu étudiés biochimiquement contrairement à leurs homologues chez E. coli.

Objectifs de l’étude :Dans le cadre de l’ITN « ProteinFactory », dont le but était d’améliorer les

capacités de production/sécrétion des protéines chez les bactéries, nous avons tenté d’identifier les

complexes membranaires impliqués dans la sécrétion de protéines chez B. subtilis en nous focalisant sur

les protéines de la voie « Sec » (Figure 1). En effet, si les connaissances fondamentales ont été obtenues

chez E. coli, peu de données existent sur les interactions physiques entre ces protéines chez B. subtilis,

ou encore leur dynamique en fonction de la protéine sécrétée ou de la phase de croissance, par exemple.

Grâce à l’emploi de deux techniques biochimiques : le BN-PAGE et l’étiquetage de protéines ciblées

pour réaliser de la purification par affinité en tandem (SPA-tag), nous avons tenté de répondre à ces

questions.

Le modèle d’étude : Bacillus subtilis est une bactérie modèle pour l’étude des bactéries à Gram positif ;

elle est capable de produire des endospores afin de résister aux conditions difficiles et est aussi un

modèle pour l’industrie par sa capacité à sécréter un grand nombre de molécules, et notamment des

protéines (Figure 2).

II

Au cours de cette étude, nous avons identifié un complexe stable formé de deux protéines membranaires,

SppA et YteJ. SppA a été décrite comme une « Signal Peptide Peptidase », une protéase qui dégrade les

peptides signaux qui s’accumulent dans la membrane cytoplasmique après libération des protéines

sécrétées. Une structure de la partie soluble de SppA de B. subtilis par cristallographie existe. Située à

l’extérieur de la cellule et ancrée à la membrane par un seul domaine transmembranaire, SppA forme

un homo-octamère [2, 3]. Son domaine C-terminal est absent dans les structures de la protéine active,

mais est présent dans un mutant catalytique, indiquant sa fonction régulatrice (Figure 3).

Par ailleurs, le gène sppA fait partie du régulon σW, qui est un facteur sigma qui devient actif lors d’un

stress affectant la membrane ou la paroi [4]. Ainsi il a été montré qu’un mutant ∆sppA est plus sensible

à la nisine (peptide antibiotique de la famille des lantibiotiques).

Enfin, le rôle prépondérant de SppA dans le recyclage des peptides signaux de la membrane a été remis

en cause par une étude qui a suggéré que RasP était la SP peptidase majeure chez B. subtilis [5]. Ceci

nous a amené à revisiter le rôle de SppA chez B. subtilis ainsi que celui de YteJ, protéine dont nous

avons découvert qu’elle formait un complexe membranaire avec SppA.

RESULTATS :

1. Identification du complexe SppA/YteJ par BN-PAGE et SPA-Tag :

1.1 Par BN-PAGE :

La technique du Blue-Native PAGE permet de réaliser une électrophorèse des complexes membranaires

dans des conditions où les interactions physiques entre les protéines sont préservées au maximum. Ainsi,

un gel de BN-PAGE d’échantillon de membranes solubilisées avec du DDM, découpé en morceaux le

long de la migration, suivi d’une analyse par spectrométrie de masse montre que les deux protéines se

III

trouvent dans la même portion de gel (Figure 4). Nous en avons conclu que SppA et YteJ co-migrent

dans ce type de gel ce qui suggère que les deux protéines forme potentiellement un complexe protéique

membranaire. YteJ, protéine inconnue, est une protéine de la famille RDD, famille de fonction inconnue,

potentiellement un transporteur.

1.2 Par SPA-tag :

La séquence d’une étiquette permettant une double chromatographie séquentielle par affinité a été

fusionnée en 5’ de la séquence du gène de la protéine d’intérêt par intégration chromosomique du

plasmide pMUTIN-SPA-LIC. Des souches sppA-SPA et yteJ-SPA furent ainsi obtenues mais seule la

souche yteJ-SPA a montré une expression de YteJ-SPA détectable par Western-blot. La purification de

YteJ-SPA à partir des membranes, solubilisées en présence de DDM, a été réalisée et les protéines co-

purifiées ont été analysées par spectrométrie de masse. La protéine la plus abondante ayant été co-

purifiée avec YteJ-SPA était SppA. Nous avons pu estimer d’après ces données de spectrométrie de

masse que le ratio entre SppA et YteJ dans le complexe était de 1:1 ou 2:1.

2. Etude des mutants de délétion de sppA et/ou yteJ :

D’après la littérature, SppA est une signal peptide peptidase, mais est aussi impliquée dans la résistance

à la nisin, un peptide antibiotique de la famille des lantibiotiques, supposément grâce à son activité

peptidase. Afin d’étudier le rôle de YteJ dans le complexe que cette protéine forme avec SppA, des

souches de B. subtilis délétées de sppA et/ou yetJ ont été construites et caractérisées.

2.1 Caractérisation des souches mutantes :

La croissance de ces souches mutantes n’est nullement affectée dans les premières 24 heures (Figure 5

A). Toutefois, la survie à 48 heures (nombre de cellules formant colonie) est particulièrement affectée

pour les souches ∆sppA et ∆sppA-yteJ, alors que la survie de la souche ∆yteJ était moins affectée. La

survie de la souche ∆sppA n’a pas pu être restaurée en co-cultivant celle-ci avec des cellules de la souche

sauvage. Il semblerait donc que l’activité de SppA soit importante sur la survie à long terme de B. subtilis.

En revanche, le phénotype observé pour la souche ∆yteJ est intermédiaire (Figure 5 B).

IV

2.2 La sécrétion des protéines :

Afin de tester le rôle de SppA et de YteJ, potentiellement via son interaction avec SppA, dans la sécrétion

des protéines, nous avons utilisé la protéine modèle AmyM dont le gène est présent sur le plasmide

pCS73. Aucune des souches comportant les délétions de sppA et/ou yteJ, n’ont montré de différence

dans la quantité finale de AmyM sécrétée dans le milieu (Figure 6). Des expériences de « pulse-

chase assays » avec ces mêmes souches mutantes réalisées par des collaborateurs (Pr JM Van Dijl,

Groningen) ont également conclu à l’absence de différence dans la cinétique de sécrétion de AmyQ.

2.3 Résistance à la subtilin

Après avoir testé la nisine, peptide de la famille des lantibiotiques, nous avons testé un autre peptide de

cette famille, la subtiline, que nous avons produite à l’aide d’une souche sécrétrice de B. subtilis

(ATCC6633). Les résultats obtenus avec la nisine et la subtiline ont été identiques, quoique plus

reproductibles avec la subtiline (Figure 7). La présence de subtiline (ou nisine) dans le milieu de culture

LB sur les cellules de la souche sauvage provoque un retard de croissance mesurable par densité optique

et ce retard est accentué chez la souche sppA à partir de 25% de subtiline. La souche yteJ montre un

phénotype intermédiaire à partir de 30%. La résistance aux lantibiotiques liée à SppA décrite dans la

littérature est donc confirmée, et nos expériences montrent que YteJ semble avoir un rôle dans cette

résistance.

A B

Figure 5 : A : Croissance en milieu LB de la souche sauvage (BSB1) et des souches délétées de sppA, yteJ

ou sppA-yteJ. B : survie à long-terme des différentes souches après comptage des cellules viables sur boites

gélosées LB-agar.

Figure 6 : Analyse par SDS-PAGE de la quantité de AmyM sécrétée dans la souche sauvage comparée aux

souches délétées de sppA, yetJ et sppA-yteJ.

V

Afin de vérifier que l’absence de SppA – et celle de YetJ - était bien la raison directe de la sensibilité

accrue à la subtiline chez les souches délétées de ces gènes, nous avons utilisé ces souches et les avons

complémentées en réintroduisant, en ectopique au locus amyE, une copie du gène délété, sppA ou yetJ,

ou encore de l’operon sppA-yteJ, sous un promoteur inductible à l’IPTG. Ces souches furent testées en

présence de 30% de subtiline, concentration retenue car provoquant une différence observable de

résistance à la subtiline entre les souches. Dans ces conditions, des concentrations croissantes d’IPTG

ont été ajouté dans le milieu LB, afin de tester la dose-réponse de l’expression de ces gènes pour la

résistance à la subtiline (Figure 8).

Figure 7 : Croissance de cellules de B. subtilis mesurée par densité optique à 600 nm dans un lecteur de

microplaques en présence de différentes concentrations de subtiline. Chaque courbe représente la moyenne

de 6 réplicats.

Figure 8 : Croissance des cellules de la souche sauvage (BSB1) et des souches délétées sppA et yetJ

complémentées de leurs gènes réciproques sous le contrôle d’un promoteur Phyperspank (Phs) inductible à l’IPTG

(notées Phs-sppA et Phs-yteJ). L’effet de la subtiline (30%) est testé en fonction de concentrations croissantes

d’IPTG (0 à 100 µM).

VI

A partir de 25 µM d’IPTG, les courbes de croissance de la souche Phs-sppA se rapproche de la souche

sauvage et de la souche Phs-yteJ, qui, elle, se comporte comme la souche sauvage. Ceci prouve que la

résistance à la subtiline dépend bien directement de SppA. En revanche, à partir de 100 µM IPTG, nous

avons systématiquement observé que les cellules de la souche Phs-sppA lysaient rapidement après

l’entrée en phase stationnaire. Il semblerait donc que la surexpression de SppA, seule, provoque un effet

délétère sur les cellules. Cet effet n’est en revanche pas observé lorsque les deux protéines SppA et YteJ

sont surexprimées en même temps comme c’est le cas de la souche Phs-sppA-yteJ (Figure 9). Nous

pouvons noter que, alors que sans IPTG cette souche est plus sensible à la subtiline, une meilleure

résistance à la subtiline pour cette souche par rapport à la souche sauvage est observable dès l’ajout de

10 µM d’IPTG dans le milieu de culture.

Ces expériences nous ont permis de confirmer le rôle direct de SppA dans la résistance à la subtiline,

possiblement via son activité protéasique/peptidasique. Cependant, une surexpression de SppA devient

délétère au-delà d’un seuil, ce qui se traduit par une lyse prématurée des cellules. En revanche, la

surexpression simultanée des deux protéines ne provoque aucun problème aux cellules de cette souche

et lui permet même d’être plus résistante que la souche sauvage.

3. Régulation de l’activité protéase de SppA La surexpression de SppA dans la souche Phs-sppA nous avait montré que, pour des concentrations de

100 µM d’IPTG, des effets délétères sur les cellules pouvaient être observés à travers une biomasse plus

faible en phase stationnaire et une chute de la densité optique certainement due à une lyse prématurée

des cellules. Cet effet était totalement annihilé par la surexpression concomitante de YteJ. Nous avons

voulu observé au microscope l’effet sur la morphologie cellulaire.

3.1 Effet de la surexpression de SppA sur la morphologie cellulaire de cellules de B. subtilis Les effets sur la morphologie cellulaire furent observés sur des cellules prises en phase

exponentielle de croissance, en présence de 100 µM d’IPTG, au microscope à contraste de phase ; les

membranes révélées avec une coloration au FM 4-64 et l’ADN chromosomique coloré au DAPI (Figure

10).

Figure 9 : Croissance des cellules de la souche sauvage (BSB1) et de la souche délétée de l’opéron sppA-

yetJ complémentés par cet opéron en ectopique sous le contrôle d’un promoteur Phyperspank (Phs) inductible à

l’IPTG (notées Phs-sppA-yteJ). L’effet de la subtiline (30%) est testé en fonction de concentrations

croissantes d’IPTG (0 à 100 µM).

VII

Pour la souche Phs-sppA, les cellules sont apparues en filament indiquant un problème de séparation des

deux cellules filles après division. La coloration au FM 4-64 a montré les septums, visibles, mais plus

éloignés les uns des autres que les septums observés pour la souche sauvage BSB1. De même la

coloration au DAPI a révélé que les chromosomes semblaient moins compacts que ceux de la souche

sauvage. Pour les souches Phs-yteJ et Phs-sppA-yteJ n’ont pas montré de différences morphologiques par

rapport à la souche sauvage. Ceci a confirmé que la surexpression de SppA, qui permet de retrouver une

résistance à la subtiline, entraine des problèmes morphologiques aux cellules et affecte le bon

déroulement du processus de division cellulaire. Nous avons vérifié que c’était bien l’activité protéase

de SppA qui en était la raison, en surexprimant une version inactive de SppA après l’introduction d’une

mutation K119A dans le site catalytique (Figure 11). Avec SppAK199A les cellules sont apparues

normales, ce qui nous a permis de conclure que c’était bien l’activité protéase de SppA qui provoquait

les défauts morphologiques.

VIII

L’effet de l’activité SppA surexprimée sur la morphologie cellulaire est annulé par la surexpression

simultanée de YteJ avec SppA dans la souche Phs-sppA-yteJ (Figure 10). Nous avons donc émis

l’hypothèse que YteJ jouait un rôle régulateur de l’activité de SppA.

3.2 Expression et purification de SppA, SppA∆1-52, du complexe SppA/YteJ et YteJ chez E.

coli.

Afin de répondre à l’hypothèse d’une régulation de l’activité de SppA par YteJ dans le complexe

SppA/YteJ identifié chez B. subtilis, nous avons exprimé chez E. coli les proteines suivantes : SppA,

SppA 1-52, SppA/YteJ (sous forme d’opéron existant nativement chez B. subtilis) et YteJ. Chaque

protéine fut exprimée avec un tag 6xHis fusionné à son extrémité N-terminale (Figure 12).

Après expression chez E. coli, excepté pour SppA∆1-52 qui a été purifiée à partir de la fraction cytosolique,

les autres protéines, étant membranaires, ont été purifiées à partir des membranes de E. coli, isolées par

ultracentrifugation et solubilisées avec 1% DDM. Les échantillons furent chargés sur colonne de Ni-

NTA agarose (Qiagen). Après lavage, les protéines furent éluées avec 200 mM imidazol et concentré

sur Amicon Ultra-15 Centrifugal Filters. L’analyse par SDS-PAGE du résultat des purifications de ces

quatre protéines est montré en Figure 13

Figure 12 : Schéma représentant les différentes expressions chez E. coli des protéines SppA, SppA1-52,

SppA/YteJ et YteJ, dont les formes sont étiquetées His en N-terminal pour une purification par colonne

d’affinité Ni-agarose.

IX

Nous avons pu conclure que SppA apparait en 2 à 3 bandes (identifiées comme SppA par spectrométrie

de masse), une seule bande pour la forme tronquée SppA∆1-52, et que la purification de His-SppA entraine

YteJ (identifiée par spectrométrie de masse) lorsqu’elle est co-exprimée avec YteJ. Ceci prouve que

l’interaction entre SppA et YteJ peut également exister chez E. coli.

3.3 Activité protéase de SppA Afin de tester l’activité protéase de SppA∆1-52, nous avons utilisé des protéines disponibles au laboratoire.

L’activité protéase de SppA∆1-52, comme annoncé dans la littérature, est apparue efficace envers

certaines protéines (HNS, Rok), partielle envers CcpA et faible voire nulle sur le lysozyme et la BSA

(Figure 14). SppA∆1-52 a montré une activité protéase dégradant seulement l’extension (linker) d’une

GFP, mais pas sur la GFP elle-même. Nous pourrions conclure que certaines structures ou séquences

d’acides aminés permettent ou non l’activité de SppA.

L’activité de SppA et de SppA/YteJ a alors été testée avec cette GFPext en tant que substrat et comparée

à celle de SppA∆1-52 (Figure 15).

A concentration de protéine identique, l’activité de SppA∆1-52 (sans domaine transmembranaire) est

beaucoup plus forte que celle de SppA (forme avec domaine transmembranaire). Cette différence peut

refléter un effet du domaine transmembranaire qui pourrait participer lui-même à la régulation de

l’activité de SppA. En revanche, l’activité de SppA dans le complexe SppA/YteJ est apparue comme

nulle durant 90 minutes. Cependant, SppA dans ce complexe est active puisque qu’une dégradation de

GFPext fut observée après 16 heures de réaction. Nous avons conclu que, dans le complexe SppA/YteJ,

YteJ agit en tant que régulateur/inhibiteur de l’activité protéase de SppA.

Figure 14 : Test de l’activité protéase de SppA∆1-52 sur différentes protéines. Après incubation de 16h, les

échantillons ont été analysés par SDS-PAGE.

X

3.4 Le domaine C-terminal de SppA des différentes formes purifiées Le domaine C-terminal de SppA est absent de SppA active mais présent dans un mutant catalytique

(K199A) ce qui suggère que ce domaine participe à la régulation de l’activité de SppA. De façon

intéressante, nous avons remarqué que la purification de SppA conduit à l’obtention de plusieurs bandes

sur gel de SDS-PAGE séparées de quelques kDa. Par spectrométrie de masse, nous avons pu identifier

que toutes ces bandes sont des formes de SppA (Figure 16). Par ailleurs, cette analyse a aussi révélé la

présence du domaine C-terminal de SppA, à travers la détection de deux peptides y appartenant,

uniquement dans SppA purifiée en complexe avec YteJ. SppA dans le complexe SppA/YteJ possède

donc encore son domaine C-terminal, comme pour le mutant catalytique SppAK199A. Ce résultat est en

accord avec le résultat précédent qui montrait que SppA dans le complexe SppA/YteJ est inactive.

3.5 Le domaine C-terminal de YteJ

Afin d’étudier plus précisément comment YteJ régule l’activité de SppA, nous avons construit des

souches qui surexpriment SppA et différentes versions tronquées du domaine C-terminal de YteJ. Des

zones structurales de YteJ ont été ciblées : dernière hélice alpha prédite (12 derniers acides aminés) ou

Figure 16 : A : Analyse par SDS-PAGE et spectrométrie de masse des différentes formes purifiées de SppA

après découpage des bandes sur gel ; B : illustration des peptides (surlignés en vert et jaune) retrouvés par

spectrométrie de masse uniquement dans la bande SppA purifiée en complexe avec YteJ.

Figure 17 : Schéma représentant les insertions de l’opéron modifié sppA-yteJ avec les différentes troncations de

yteJ. Ces constructions ont été insérées dans le chromosome de B. subtilis au locus amyE.

XI

encore le dernier domaine transmembranaire potentiel (35 derniers acides aminés) (Figure 17). Pour

tester l’effet de ces troncations de YteJ sur l’activité SppA, nous nous sommes basés sur le fait que

l’activité SppA est négativement régulée par YteJ et que, si SppA est non-régulée donc active, alors in

vivo les cellules apparaissent allongées au microscope.

Les cellules furent observées au microscope en phase exponentielle de croissance dans un milieu LB

avec 0 ou 100 µM IPTG. Les cellules surexprimant SppA et les troncations de YteJ montrent que

l’activité de SppA est dérégulée dès que YteJ est tronquée de ses 12 derniers acides aminés (cellules

allongées), et encore plus avec la troncation des 35 derniers acides aminés (Figure 18). Nous pouvons

conclure que le domaine C-terminal de YteJ contient des déterminants qui sont importants dans le

contrôle de l’activité de SppA.

CONCLUSION et PERSPECTIVES :

Chez B. subtilis, la protéase SppA forme un complexe membranaire avec YteJ. L’implication de SppA

dans la maturation des protéines sécrétées n’a pas pu être mis en évidence, mais nous avons confirmé

son rôle direct dans la résistance aux peptides antibiotiques de la famille des lantibiotiques. Le rôle de

YteJ a été analysé et nous avons montré, in vivo et in vitro, que YteJ régule négativement l’activité de

SppA. Cette activité de SppA, lorsque surexprimée, apparait délétère pour la cellule. La surexpression

simultanée de YteJ abolit cet effet délétère, renforçant l’idée que YteJ régule l’activité de SppA. Plus

précisément c’est le domaine C-terminal de YteJ qui semble être impliqué dans cette régulation. Afin

de mieux comprendre cette régulation, la topologie du domaine C-terminal de YteJ devrait être étudié

ainsi que les déterminants majeurs de son interaction avec SppA. Une caractérisation du complexe dans

son ensemble par microscopie cryo-microscopie électronique apporterait des réponses sur son

organisation générale dans la membrane. Physiologiquement, il reste encore une question importante :

si YteJ est bien capable d’inhiber SppA, quels sont les signaux extérieurs (comme les lantibiotiques) qui

entraine l’activation de SppA, c’est-à-dire la levée de l’inhibition par YteJ. Et via quel mécanisme

moléculaire cette levée d’inhibition peut-elle se faire au niveau de YteJ et SppA. Une étude récente sur

le gène rdd chez Halobacillus andaensis, potentiellement homologue de yteJ chez B. subtilis, ouvre des

pistes sur une fonction d’antiporteur Na+/H+ si celle-ci a été conservée entre ces deux espèces [6].

REFERENCES:

1. Tjalsma, H., et al., Proteomics of protein secretion by Bacillus subtilis: separating the

"secrets" of the secretome. Microbiol Mol Biol Rev, 2004. 68(2): p. 207-33.

2. Nam, S.E., A.C. Kim, and M. Paetzel, Crystal structure of Bacillus subtilis signal peptide

peptidase A. Journal of Molecular Biology, 2012. 419(5): p. 347-358.

FM 4-64 DAPI Merge

IPTG

Figure 18 : Observation au microscope à contraste de phase et fluorescence de l’état morphologique des cellules

surexprimant SppA et différentes troncations de YteJ, en absence d’IPTG (1ère colonne) et en présence de 100

µM d’IPTG (4 dernières colonnes) dans le milieu de culture.

XII

3. Nam, S.-e. and M. Paetzel, Structure of Signal Peptide Peptidase A with C ‑ Termini Bound

in the Active Sites: Insights into Speci fi city, Self-Processing, and Regulation. Biochemistry,

2013. 52: p. 8811-8822.

4. Kingston, A.W., X. Liao, and J.D. Helmann, Contributions of the σ(W) , σ(M) and σ(X)

regulons to the lantibiotic resistome of Bacillus subtilis. Molecular microbiology, 2013. 90(3):

p. 502-18.

5. Saito, A., et al., Post-liberation cleavage of signal peptides is catalyzed by the site-2

protease (S2P) in bacteria. Proceedings of the National Academy of Sciences of the United

States of America, 2011. 108(33): p. 13740-5.

6. Shao, L., et al., Characterization of a Functionally Unknown Arginine–Aspartate–Aspartate

Family Protein From Halobacillus andaensis and Functional Analysis of Its Conserved

Arginine/Aspartate Residues. Frontiers in Microbiology, 2018. 9: p. 807-807.

Acknowledgments

This study was carry out at the Systems Biology for bacterial Engineering and Redesign

(SyBER) group, part of the Genetic Controls in Bacterial Systems (GCBS) group, at the Microbiologie

de l’Alimentation au service de la Santé (MICALIS) Institute, INRA, Jouy en Josas (France). This PhD

project was part of the European Marie Skłodowska-Curie ITN programme: ProteinFactory -

"Engineering of New-Generation Protein Secretion Systems''. The development of an international PhD

project it is not an easy road but with the support, encouragement and generosity of a large number of

people, including my family, friends, colleagues and collaborators it has been continued towards its

completion. All of them deserve special appreciations.

Je tiens à exprimer mes remerciements aux membres du jury, qui ont accepté d’évaluer mon

travail de thèse: Anne Galinier et Nathalie Declerck pour avoir accepté d’être les

rapporteurs de mon manuscrit. Je remercie également Sylvie Nessler, Nathalie Campo, Martin Picard et

Jean Luc Pernodet d’avoir accepté d’examiner ce travail. Je leur suis reconnaissante pour toutes les

corrections et suggestions qu’ils apporteront.

First of all, I would like to start by thanking to my supervisor Olivier Delumeau for accepting

me as PhD student, welcoming me to the team and for helping me to settle down on my arrival. I greatly

appreciate Olivier for what he taught me during my PhD thesis, not only about the scientific knowledge

he transferred to me, but also the importance of transferable skills as resilience. I would like to express

my profound gratitude to our PI, Matthieu Jules, for his genuine leadership, commitment, support and

encouragement. He believed and motivated me when I most needed, saying the right words at the right

time, as well as pushed me to give my best on the most difficult times. I am grateful for having the

opportunity of working and learning from him. To my bench mate, Stephen McGovern, I will be always

grateful for all what he taught me, helped me and for all our conversations and jokes! I would like to

thank to the rest of our team members for all the nice moments we passed together, great discussions

and shared ideas. Specially to Etinne Dervyn, for all our great conversations and for your advices, to

Claire and Marwa (my Protein factory roommate) for all we have shared and support each other along

the way. Je vous remercie tous de faire partie de ce voyage!

When you live abroad, far from your home town, friends start to be like a family you have there.

I am grateful for the friends I have made these years and for all the great moments we have shared, you

are one of the best things I bring from this experience! Aos melhores vizinhos que podia pedir, obrigado

por tudo Joana, João e Lina! Pela cumplicidade imediata e por toda a amizade que criamos, fica o meu

obrigado do fundo coração Priscilla, Caroline e Camilla, que mesmo transatlântica que se mantenha! À

minha companheira de zumba Transito e à minha italiana favorita Ludovica gracias por toda a amizade,

força e jantares temáticos! To ma coupine and my sweetie Cecile, I have no words to express what I feel

about all the moments we have shared. Thank you my sweetie, we were together in the life ups and

downs, and our craziness!

To the amazing experience that was Protein Factory consortium! It was a privilege work and

learn with this amazing group of talented people. Thank you for all the knowledge shared and for all the

traveling we did together! First I would like to express my gratitude to Jan Maarten van Dijl, for joining

together this consortium, for being an example of true leadership, support and commitment. Thank you,

Jan Maarten van Dijl and Colin Robinson, despite all the tiredness, for stay up late with us after meetings,

for drinks and advices. To Sierd Bron, thank you for helping us and for sharing with us your passion

about science. To all the members of my thesis committee (Jan Maarten van Dijl, Emma Dave, Edda

Klipp, Andreas Otto and Tjeerd van Rij) for all the productive discussions and guidance. I would like to

express my gratitude to David Humphreys, Emma Dave, Neesha Dedi and Gilles Malherbe for

welcoming me to UCB Celltech and for all the help during my secondment. Also to Dörte Becher,

Andreas Otto and Minia Varela for welcoming me to Greifswald and helping me during my secondment.

Os três anos do meu doutoramento foi mais uma prova da quão sortuda sou por ter uma família

e amigos maravilhosos, que independentemente da distância ou quaisquer outros fatores, estiveram

sempre lá para me apoiar! Sou muito grata por isso!

Grata pelas minhas tias e primos, que me ensinam todos os dias a ser o melhor de mim, sendo

exemplos de força e amor, e que estiveram sempre lá para o bom e mau, para me dar colo quando

precisei. Ao meu primo-irmão Alexandre, por tudo o que vivemos juntos, obrigado (e também por me

ensinares as primeiras asneiras em francês deu jeito)! Um enorme obrigado ao meu pai e à Irene pela

ajuda, apoio, pelo carinho e as palavras certas no momento certo, que me deram força quando mais

precisei! Um obrigado aos meus Sogros, ao Hugo e Daniel por também fazerem parte desta jornada,

partilharem comigo bons e maus momentos, e por todo o carinho!

Não tenho palavras para expressar a minha gratidão, pelas minhas companheiras de vida, de

sempre e para sempre, Sofia, Joana e Patrícia! Muito obrigado por estarem sempre presentes! E claro ao

resto da nossa família, claro a que escolhi (sei eu que estou a ser lamechas) João, Filipe e Marisa!

À pessoa que estarei eternamente grata, a minha Mãe, não só porque sempre fez de tudo para

me ajudar a ser a pessoa que sou hoje como é uma guerreira cheia de força. Foi ela que me ensinou o

valor da vida, da família e a ir sempre á luta pelos meus sonhos! Obrigado Mãe!

Como os ingleses dizem, for last and not least, o meu amor para a vida toda, o Diogo! Esta tese

também é tua, sem dúvida que sem ti isto não teria sido possível! Estiveste sempre ao meu lado e muitas

vezes confiaste mais em mim do que eu própria e isso nunca vou saber como te agradecer o suficiente!

Os estudos diziam que a maioria dos relacionamentos não sobrevivem a uma tese de doutoramento, nós

mais uma vez contra estatísticas viemos mostrar que o amor cura, dá força e vence tudo! Obrigado

Diogo!

Queria por último expressar a minha gratidão por o meu caminho se ter cruzado com a lendária

equipa da becoach! A grande turma do Arranque90 que no momento certo veio me ajudar a olhar para

a minha vida de um outro prisma! Com especial agradecimento ao Marco Fonseca e ao Thiago Feder,

pela oportunidade que me deram, pela ajuda e força, para que seja a melhor versão de mim!

“Life doesn’t give you the people you want. It gives you the people you need: to

love you, to help you, to hurt you, to leave you and to make you the person you

were meant to be!”

To all of you that believed in me!

“We must have perseverance and above all confidence in ourselves. We

must believe that we are gifted for something and that this thing must

be attained.”

Marie Curie

LIST OF ABBREVIATIONS

ATP – Adenosine triphosphate

ADP – Adenosine diphosphate

CaM – Calmodulin resin

Cm – Chloramphenicol

DDM – n-Dodecyl β-D-maltoside

EDTA – Ethylenediaminetetraacetic Acid

EGTA – Egtazic Acid

Ery – Erythromycin

FEA – Flagella Export Apparatus

ICE – Integrative and Conjugative Elements

IPTG – Isopropyl β- d-1-thiogalactopyranoside

GFP – Green Fluorescent Protein

GRAS – Generally Recognized as Safe

LCA – Live-Cell Array

LIC – Ligation-independent cloning

LTA – Lipoteichoic Acid

Neo – Neomycin

ORF – Open Reading Frame

PMF – Proton Motor Force

RBS – Ribosomal Binding Site

RNC – Ribosome-Nascent Chain

SP – Signal Peptide

SPA – Sequential Peptide Affinity

SPase – Signal Peptide Peptidase

SppA – Signal Peptide Peptidase A

Spec – Spectinomycin

SRP – Signal Recognition Particle

TCA – Trichloroacetic Acid

T4SS – Type IV Secretion System

WTA – Wall Teichoic Acid

1

TABLE OF CONTENTS

3

TABLE OF CONTENT

1 INTRODUCTION ............................................................................................................................. 13

1.1 General introduction ............................................................................................................. 13

1.1.1 Project context .............................................................................................................. 13

1.1.2 Bacillus subtilis, a Gram-positive bacterium ................................................................. 14

1.1.3 B. subtilis: a Gram-positive model ................................................................................. 16

1.1.4 B. subtilis: a bacterium of industrial interest ................................................................ 17

1.2 Protein translocation across cytoplasmic membrane in bacteria ......................................... 19

1.2.1 The general protein secretion pathway: the Sec system .............................................. 20

1.2.2 Substrate recognition .................................................................................................... 22

1.2.3 Intracellular chaperoning and targeting to the Sec translocase ................................... 25

1.2.4 Twin-Arginine Transport (Tat) Pathway ........................................................................ 33

1.2.5 Flagella Export Apparatus (FEA) .................................................................................... 38

1.2.6 Type IV Secretion Systems ............................................................................................. 43

1.2.7 Holins ............................................................................................................................. 47

1.3 A guarantee of a correct protein translocation in B. subtilis ................................................ 49

1.3.1 A bacterial cytoplasmic quality control system: ClpXP protease .................................. 49

1.3.2 Membrane-bound proteases, an important quality control step for membrane and

secretory proteins. ........................................................................................................................ 54

1.3.3 Signal peptide peptidase A (SppA) ................................................................................ 55

1.4 Scope of the Thesis ................................................................................................................ 60

2 Results ........................................................................................................................................... 65

2.1 Identification of membrane protein complexes involved in the B. subtilis Sec pathway ..... 65

2.1.1 SPA-tag constructions .................................................................................................... 65

2.1.2 Purifications of the SPA-tagged proteins ...................................................................... 69

2.1.3 Optimisation of the BN-PAGE technique ....................................................................... 71

2.2 The SppA/YteJ membrane complex ...................................................................................... 74

2.2.1 The sppA yteJ operon is regulated by sigma factors σA and σW ..................................... 74

2.2.2 Deletion of the sppA and yteJ genes ............................................................................. 80

2.2.3 Role of the SppA/YteJ complex in protein secretion ..................................................... 82

2.2.4 The deletion of sppA yteJ affects the long term survival .............................................. 87

2.2.5 The deletion of sppA and yteJ affects the resistance to lantibiotics ............................. 90

2.2.6 The deletions of sppA and yteJ do not affect the resistance to vancomycin,

erythromycin or protect against lysozyme .................................................................................... 94

2.2.7 Overexpression of sppA results in the recovery of the BSB1 phenotype in the presence

of subtilin. ...................................................................................................................................... 97

4

2.2.8 The overexpression of sppA results in a change of the cell morphology. ................... 100

2.2.9 SppA and YteJ cell localization..................................................................................... 103

2.2.10 SppA and YteJ in vitro purification. ............................................................................. 104

2.2.11 SppA can digest fully folded proteins or not ............................................................... 106

2.2.12 YteJ inhibits SppA activity ............................................................................................ 107

2.2.13 The C-terminal domain of YteJ is involved in the regulation of SppA activity ............ 110

2.2.14 SppA digests subtilin .................................................................................................... 112

3 Discussion and perspectives ........................................................................................................ 117

3.1 Protein interactions of the B. subtilis Sec pathway ............................................................. 117

3.2 SppA/YteJ membrane protein complex .............................................................................. 118

3.2.1 Role of SppA and YteJ in protein secretion: ................................................................ 119

3.2.2 Role in the resistance to nisin, subtilin and LP9 .......................................................... 120

3.2.3 Regulation of the sppA yteJ operon ............................................................................ 120

3.2.4 SppA/YteJ, a complex involved in quality control of cell division? ............................. 121

3.2.5 The protease activity of SppA and its regulation by YteJ ............................................ 123

3.2.6 YteJ, another function? ............................................................................................... 125

3.2.7 Model for the interaction of SppA and YteJ ................................................................ 126

3.3 Main results ......................................................................................................................... 127

4 MATERIAL AND METHODS .......................................................................................................... 131

4.1 Techniques for DNA manipulation ...................................................................................... 131

4.1.1 Oligonucleotide ........................................................................................................... 131

4.1.2 Polymerase chain reaction (PCR) ................................................................................ 131

4.1.3 Purification of PCR products ........................................................................................ 131

4.1.4 Electrophoresis of DNA ............................................................................................... 131

4.1.5 Purification of DNA from agarose gel .......................................................................... 131

4.1.6 Purification of plasmid DNA ........................................................................................ 132

4.1.7 Chromosomal DNA extraction ..................................................................................... 132

4.1.8 Digestion of DNA with restriction enzymes ................................................................ 132

4.1.9 Ligation of DNA fragments .......................................................................................... 132

4.1.10 Gibson assembly technology ....................................................................................... 132

4.1.11 Ligation-Independent Cloning (LIC) ............................................................................. 133

4.1.12 DNA sequencing .......................................................................................................... 135

4.1.13 DNA transformation .................................................................................................... 135

4.2 Bacterial strains and growth conditions .............................................................................. 135

4.3 Strain construction .............................................................................................................. 136

4.3.1 SPA tagged strains construction .................................................................................. 136

5

4.3.2 Deletion mutants ......................................................................................................... 138

4.3.3 SppA, YteJ and SppA-YteJ overexpressing strains ....................................................... 139

4.3.4 Construction of the GFP Fusions ................................................................................. 140

4.3.5 His- tag constructions .................................................................................................. 140

4.4 Fluorescence Microscopy .................................................................................................... 141

4.5 Live-cell array (LCA) ............................................................................................................. 141

4.6 Bacteria survival test ........................................................................................................... 141

4.7 Swimming and swarming capacity ...................................................................................... 142

4.8 Biofilm formation ................................................................................................................ 142

4.9 Protein analysis ................................................................................................................... 142

4.9.1 Sodium Dodecyl Sulphate Poly-Acrylamide Gel Electrophoresis (SDS PAGE gels) ...... 142

4.9.2 Blue Native PAGE ......................................................................................................... 143

4.9.3 Western blot analysis .................................................................................................. 143

4.9.4 Secreted proteins analysis ........................................................................................... 144

4.9.5 Sequential Peptide Affinity (SPA) purification ............................................................. 144

4.9.6 His-tag purification ...................................................................................................... 146

4.9.7 Double tag purification ................................................................................................ 147

5 BIBLIOGRAPHY ............................................................................................................................. 151

6 Appendix ...................................................................................................................................... 179

6.1 Contents, sterilization and storage of required solutions ................................................... 179

6.2 UCB Secondment: Purification and characterization of a maltogenic α-amylase from

Bacillus stearothermophilus ............................................................................................................ 198

7

LIST OF FIGURES

Figure 1. Schematic representation of the distinct cell fates in Bacillus subtilis biofilms…………....16

Figure 2. Protein export pathways in B. subtilis………………………………...………………….....19

Figure 3. Schematic prediction of the B. subtilis Sec-dependent secretory protein translocase……....21

Figure 4. Components involved in Sec-dependent protein export in B. subtilis………………...…….22

Figure 5. General features of the signal peptides and propeptides of Bacillus secretory proteins…....24

Figure 6. Schematic representation of the different modes of protein translocation across the

membrane of B. subtilis……………………………………………………………………...26

Figure 7. Scheme of SecA-mediated protein translocation………..……………………………….....29

Figure 8. Signal peptide cleavage of precursor proteins by signal peptidases….…….……………….31

Figure 9. Tat pathway components………………………………………………………………...….34

Figure 10. Gram-positive bacterial Tat pathway..…………………………………………………….35

Figure 11. Model of the regulation of TatA pore formation by TatBC and substrate……..………….36

Figure 12. Bacterial flagella structure………………………………………………………………....39

Figure 13. B. subtilis flagellum…..……………………………………………………………….…...40

Figure 14. Model for the assembly process of the Gram-negative flagellar type III export apparatus 41

Figure 15. An entropic chain mechanism for flagellum growth outside the cell………….…….…….42

Figure 16. Schematic representation of type IV secretion architecture and functions in bacteria….....44

Figure 17. T4SS of ICEBs1……………………………………….………………………..………....46

Figure 18. Schematic depiction of the proteins involved in Gram-negative bacterial cell envelope

disruption by holin-type lysis systems………………………………………………………48

Figure 19. Scanning electron micrographs……………………………………………………………50

Figure 20. Cartoon model of substrate recognition and degradation by the ClpXP protease….….…..51

Figure 21. Interaction of ClpX and ClpP……………………………………………………………...51

Figure 22. Representation of the expansion and contraction of the axial pore by a snake-jaws

model………………………………………………………………………………………...52

Figure 23. The B. subtilis SppA protein………………………………………….…………………...56

Figure 24. Comparison between octameric SppABS and tetrameric SppAEC…………….………....57

Figure 25. Cross section representation of the oligomeric dome shape structure of SppAEC and

SppABS………………………………………………………………………………….…..58

Figure 26. Growth analysis of strains harbouring C-terminal SPA-tag fusions………………..……..67

Figure 27. Expression of the SPA tag fused proteins…………………………………..….………….68

Figure 28. Purification of SpoIIIJ-SPA and SecDF-SPA protein with different concentrations of

DDM………………….………………………………………………………….……….….70

Figure 29. Analysis by Western blot of a Blue native PAGE 4-16% Bis-Tris Gel of SecDF-SPA in the

presence of different concentrations of DDM………………………………….……………72

8

Figure 30. Number of peptides identified, by LC-MS/MS for each of the proteins purified in the

presence of 1% DDM………………………………………………………………………..73

Figure 31. Alignment of sppA and yteJ genes with other bacteria of firmicutes phylum……..…..…..74

Figure 32. Transcriptional fusions of the PσAW and PσW, promoters and sfGFP………………...……75

Figure 33. Nisin induced increasing expression of sfGFP, regulated by σA and σW sigma factors…...77

Figure 34. Subtilin induced increasing expression of sfGFP, regulated by σA σW sigma factors……..79

Figure 35. Growth of B.subtilis sppA and yteJ deletion mutant strains……………………………….81

Figure 36. Cell morphology of the sppA yteJ deletion mutant……………………………..…………82

Figure 37. Analysis of AmyM extracellular amount by SDS PAGE………………………..………..84

Figure 38. Analysis of AmyM extracellular amount by Western Blot………………………..………85

Figure 39. SDS PAGE analysis of AmyM extracellular amount produced by B. subtilis WT and

overexpression mutant strains……………………………………………………………….86

Figure 40. Capacity of survival of BSB1 and ∆yteJ, ∆sppA and ∆sppA yteJ deletion mutants...…….88

Figure 41. Long-term survival of BSB1 and ∆sppA yteJ deletion mutant, in LB and M9 liquid

media.......................................................................................................................................89

Figure 42. Complementation assays in LB with BSB1 and ∆sppA yteJ deletion mutant strains……..90

Figure 43. The growth of B. subtilis sppA and yteJ deletion mutant strains in the presence of different

concentrations of Nisin……………………………………………………...…………….…91

Figure 44. Growth of the B. subtilis sppA and yteJ mutant strains in the presence of different

concentrations of subtilin…………………………………………………….……………...92

Figure 45. The growth rates of BSB1, ∆sppA and ∆yteJ in the presence of different concentrations of

subtilin………….……………………………………………………………………………94


concentrations of vancomycin………………………………………………………………95


concentrations of erythromycin……………………………………………………………..96

Figure 48. The B. subtilis sppA and yteJ deletion mutant strains treated with different concentrations

of lysozyme…………………………….................................................……………………97

Figure 49. The B. subtilis sppA and yteJ overexpressing strains……………………………...………99

Figure 50. The B. subtilis sppA yteJ overexpressing strain…………………..……..……………….100

Figure 51. Cell morphology of the sppA, yteJ or sppA yteJ overexpression mutants……….………102

Figure 52. Cell morphology of the sppA and sppAK199A overexpression mutants………..…….…….103

Figure 53. SppA and YteJ cell localization………………………………………………………….104

Figure 54. Scheme representing the different His-tag constructions………………..……………….105

Figure 55. SDS PAGE analysis of SppA∆1-52, SppA, SppA/YteJ and YteJ His-tagged purified

proteins……………………………………………………………………………………..106

Figure 56. SDS PAGE analysis of SppA∆1-52 activity………………………………………………..107

9

Figure 57. SDS PAGE analysis of SppA∆1-52, SppA and SppA/YteJ activity……………………….109

Figure 58. SDS-PAGE and mass spectrometry analysis of SppA and SppA/YteJ complex…...……109

Figure 59. Scheme of SppA/YteJ protein complex in the different YteJ truncated constructions…..110

Figure 60. Cell morphology of the overexpressing strains…………………….……………….……111

Figure 61. Growth of BSB1 and ∆sppA in the presence of different subtilin solutions……….…….113

Figure 62. Cells mutated for minJ are filamentous……………….……………………………….…122

Figure 63. Morphology of B. subtilis cells lacking multiple penicillin-binding proteins (PBP)…….122

Figure 64. Amino acid sequence alignment between H. andaensis RDD and B. subtilis YteJ…..….125

Figure 65. Model for the regulation of SppA by YteJ………………………………..……………...126

Figure 66. LIC integration into B. subtilis…………………………………...………………………136

Figure 67. Sequential peptide affinity purification……..…………………….……………………...146

LIST OF TABLES

Table 1. Set of proteins involved in the Sec pathway chosen to be SPA-tagged for tandem affinity

purification……………………………….…………………..……………………………...65

Table 2. Oligonucleotides used for LIC constructions………………………………………...……..137

LIST OF APPENDIX TABLES

Table S 1. Stock and working solutions of growth supplements…………………..………………...180

Table S 2. Stock and working solutions of antibiotics…………………………...…………………..180

Table S 3. List oligonucleotides………………………………………………………...……………181

Table S 4. List of B. subtilis strains………………….………….…………………………………...188

Table S 5 List of E. coli strains……………………………………………………………...……….190

Table S 6 List of plasmids………………………………………………………..….……………….191

LIST OF APPENDIX FIGURES

Figure S 1 YteJ predicted structure by Protter…………………………………...…………………..193

Figure S 2. Purification of SppA, YteJ, SppA/YteJ and SppAK199A by Gel Filtration…………….....194

Figure S 3 Pulse chase assays for sppA, yteJ and sppA yteJ deletion mutants and complemented

strains……………………………………………………………………………………….195

Figure S 4. The growth of B. subtilis sppA and yteJ deletion mutant strains in the presence of different

concentrations of LP9………………………………………………………………...…….196

Figure S 5. The growth of B. subtilis sppA and yteJ overexpressing mutant strains in the presence of

different concentrations of LP9…………………………………………………………….197

Figure S 6. AmyM supernatant purification…………………………………...…………………….199

Figure S 7. AmyM Size exclusion purification results……………………………………...……….200

11

INTRODUCTION

Chapter 1. INTRODUCTION

13

1 INTRODUCTION

1.1 General introduction

1.1.1 Project context

Biotechnology has become one of the most important industries in our modern society. This

industry promotes the development of technology and products based on the use of living organisms

and their products for the benefit of mankind in different areas and, at the same time, contributes to a

resource-efficient and sustainable economy (Gavanji 2013; Chu and Robinson 2001). The biotech

industry is hugely important for the EU, as it is a sector that employs around 7.5 million people and it

has an annual turnover over 100 € billion (European Commission 2017). Within this industry, the

production of recombinant enzymes and biopharmaceutical proteins is of major importance. The

global market for such recombinant proteins has grown significantly in the past decade, worth 180 €

millions in 2017 and is forecast to exceed 225 € million by 2021. The constant growth of this market

leads to a continuous need for optimization of the production platforms that can deliver protein

products in greater yields, with higher quality and at lower costs. Therefore there is an overwhelming

need for the development of next-generation super-secreting cell factories (van Dijl and Hecker 2013;

Browning et al. 2017; Matthews et al. 2017).

This thesis was carried out within the frame of the EU-funded innovative training network

(ITN), Protein Factory (2015-2018), of the Horizon 2020 Marie Skłodowska-Curie actions program.

This project brings together 6 academic Institutes, which provide the expertise in synthetic biology

and protein secretion, and 5 non-academic partners which include some of the world's premier

biotechnology companies (Novozymes, DSM, Abera Bioscience, UCB-Celltech and FGen) with the

main objective of creating super-secreting strains, bypassing major production bottlenecks such as

secretion stress. A variety of different host expression systems for the production of recombinant

proteins has been described over the years, and each has its own advantages and disadvantages. The

host organisms should be precisely selected for each specific product, considering the product

development and running cost, yield, and easiness of downstream processing, time to market,

necessity of correct glycosylation, and available infrastructure (Chu and Robinson 2001). The two

most popular industrial workhorses for the production of biopharmaceuticals and industrial enzymes

are Escherichia coli and Bacillus subtilis, respectively (Tu, Hong, and Chen 2009). These bacteria are

the two model organisms used in Protein Factory. Both are fast growing bacteria able to replicate in

cheap media to high cell density, their genomes are far better studied than other microorganisms which

makes available many genetic tools, both are amenable to genetic manipulation and can express and

produce proteins with high yields. Due to E. coli very high level of expression of recombinant

proteins, that riches up to 30% of total cellular protein, this bacterium is preferably used for the


14

production of biopharmaceuticals, such as the human growth hormone (e.g. Humatrope) or insulin

(e.g. Lispro) (Ramadan et al. 2015). E. coli is a Gram-negative bacterium, characteristically having a

periplasm and an outer membrane bilayer composed of lipopolysaccharides. As such, the secreted

proteins are exported from the cytoplasm to the periplasmic space from which they can be recovered

by the rupture of the outer membrane, which gives advantages in downstream processing (Blount

2015). However, the presence of this outer membrane can be a limitation as it works as a permeability

barrier and obstructs the secreted proteins of being released to the medium. The overexpression of the

recombinant proteins in this host can origin misfolded proteins that accumulate in the cytoplasm as

inclusion bodies (Villaverde and Carrió 2003). To recover these proteins the inclusion bodies need to

be solubilized and the proteins refolded into bioactive molecules, a process that bring major costs to

the production process with a final poor recovery of the recombinant proteins (Sørensen and

Mortensen 2005). At an industrial scale and depending on the proteins to be obtained, in the case of

secreted proteins, using B. subtilis as a host organism might be a better option. As a non-pathogenic

Gram-positive bacteria, B. subtilis have a cytoplasmic membrane surrounded by a permeable cell

envelope that allows the release of secretory proteins directly into the culture medium. Therefore, B.

subtilis was used as a model organism for the development of this thesis project and, will be described

in detail in the next section.

1.1.2 Bacillus subtilis, a Gram-positive bacterium

Bacillus subtilis is a Gram-positive rod-shaped bacterium that belongs to the Firmicute

phylum (Bacillaceae family). The main classes of this phylum are the Clostridia, Bacilli (Bacillus,

Listeria, Enterococcus, Staphylococcus and Streptococcus) and Mollicute (Wolf et al. 2004). B.

subtilis was discovered in 1835 by Christian Gottfried Ehrenberg and originally named “Vibrio

subtilis”, however in 1872 Ferdinand Cohn renamed it “Bacillus subtilis”. B. subtilis is considered as

an obligate aerobe, but can also grow anaerobically by respiration on nitrate or by fermentation on

glucose (Nakano et al. 1997; Hoffmann et al. 1995). The B. subtilis genome exhibits a low % GC

content and, although Gram-positive and Gram-negative bacteria diverged about one billion years ago,

it shares similarities with the E. coli genome sequence, as about one-quarter of the entire B. subtilis

Box1: Project context

This thesis is part of the EU-funded innovative training network (ITN), part of the

Horizon 2020 Marie Skłodowska-Curie actions program, entitled Protein Factory. A project

that intends to overcome the major bottlenecks of recombinant protein production, such as

secretion stress, through the creation of engineered super-secreting strains, capable of

secreting an unprecedented range of target molecules. Bacillus subtilis was the model

organisms used in this project.


15

genome have clear orthologous counterparts in E. coli (Kunst et al. 1997). Besides the core of the

translation and transcription machineries, other operons involved in major integrated functions, such

as ATP synthesis (atp operon) and electron transfer machinery (cta and qox operons), are well

conserved between the two organisms (Kunst et al. 1997). In contrast, these two bacteria are

structurally different, B. subtilis, as a Gram-positive bacterium, possesses a single cytoplasmic

membrane and a thick cell wall matrix of peptidoglycans and teichoic acids, thus lacking a true

periplasmic space (Matias and Beveridge 2005; Christopher Weidenmaier and Peschel 2008), while E.

coli has a characteristic Gram-negative envelope composed by three principal layers, the inner (IM)

and outer (OM) membrane flanking their cell wall within the periplasmic space (Salton 1953;

Christopher Weidenmaier and Peschel 2008).

B. subtilis natural niche is the rhizosphere soil, it grows associated to the plant roots, a healthy

and mutually beneficial association. By one side, B. subtilis benefits from the components excreted by

plant roots (such as amino acids, organic acids, sugars, phenolic acids and other secondary

metabolites), in turn it contributes to the plant defence against pathogenic bacteria (Gaskins, Albrecht,

and Hubbell 1985; Haney et al. 2015).

B. subtilis is able to survive in or adapt to adverse conditions, to engage various life-styles by

triggering different genetic programs, in response to unfavourable environmental signals (Mielich-

Süss and Lopez 2015; López and Kolter 2010). B. subtilis responds to different environmental changes

or to the lack of an essential nutrient, by differentiating into subpopulations of specialized cells:

I. motile cells, give the capacity to the cells to move through liquid and over surfaces;

II. endospore, a small robust, enduring and metabolically inactive cells are produce in

response to nutritional limitations. Spores can survive years in the absence of

nutrients, and can resist to a wide variety of extreme conditions such as heat, cold,

drying and radiations (Burbulys, Trach, and Hoch 1991; Gonzalez-Pastor 2003).

During the sporulation process, cells secrete extracellular killing factors that induce

the lysis of nonsporulating cells that have not developed immunity to these toxins.

This lysing process releases nutrients into the medium that the surviving sporulating

cells can feed on, and thus this behaviour is referred to cannibalism (González-Pastor

2011; Gonzalez-Pastor 2003);

III. competent cells, a type of cells able to scavenge DNA from environment to use as a

nutrient source or, when possible, integrated into the chromosome by recombination

to gain new functions (Anagnostopoulos and Spizizen 1961);

Connected to the cell fate is their social behaviour, bacteria do not grow as dispersed

individual organisms, they communicate and cooperate with each other, originating bacterial

communities. This social behaviour gives to the cell the ability to perceive environmental changes and

react accordingly (López and Kolter 2010). The biofilms are an example of how bacterial cells can


16

form multicellular structures able to adhere to surface encased in a self-produced extracellular matrix,

as it is visible on Figure 1 (Vlamakis et al. 2013; Kearns et al. 2005; Aleti et al. 2016).

Undomesticated B. subtilis strains form robust, multicellular biofilms that exhibit complex

architectural features (Lemon et al. 2008). Swarming is another social behaviour through which a

synchronised group of flagellum-driven bacterial cells moves on a surface (Partridge and Harshey

2013; Kearns and Losick 2003; Ke et al. 2015). Bacteria communicate between each other and

respond to fluctuations in external cues as a global bacterial population by a complex sensing system

called quorum sensing. This mechanism allows the cells to produce signal molecules to activate

distinct pathways and trigger changes in gene expression in response to changes in cell density,

leading bacteria to adapt to new scenarios (Abisado et al. 2018; Camilli and Bassler 2006; Ke et al.

2015).

Figure 1. Schematic representation of the distinct cell fates in Bacillus subtilis biofilms. A mature biofilm is

composed of distinct subpopulations of cell types that coexist and exhibit different spatiotemporal distribution

patterns. The different subpopulations interact and cooperate with each other with the objective of overcome

changes of external and environmental factors. Adapted from Mielich-Süss and Lopez 2015.

1.1.3 B. subtilis: a Gram-positive model

In 1958, John Spizizen showed for the first time that the strain B. subtilis 168, a non-

pathogenic and auxotrophic for tryptophan strain, is transformable by exogenous chromosomal DNA

(Spizizen 1958). This discovery opened a door that allowed the scientific community to study and

deeply understand the genetics and molecular biology of B. subtilis, adopting this organism as a

genetic tool, which is one of the most studied microorganisms along with E. coli and Salmonella

typhimurium. The 4215 kb entire genome of B. subtilis was sequenced in 1997 and completely re-


17

sequenced and annotated in 2009, with 4406 predicted ORFs (Open Reading Frame) (Kunst et al.

1997; Barbe et al. 2009). Studies showed that only around 260 genes were found to be essential for

growth, corresponding to proteins involved in DNA replication, protein synthesis (i.e. ribosomes,

synthesis, translation, secretion, protein quality control, lipid biosynthesis, cell metabolism and cell

division) (Kobayashi et al. 2003; Commichau, Pietack, and Stülke 2013; Koo et al. 2017). The

acquirement of such information, as the genome sequence, has allowed more comprehensive and

global experimental approaches such as transcriptomic, proteomics and interactomics (Mäder et al.

2002; Otto et al. 2010; Marchadier et al. 2011; Nicolas et al. 2012). Along with the creation of

dedicated and reliable B. subtilis databases. These databases are either part of global efforts such as

BsubCyc or specialized to specific scientific problems such as transcriptional regulation in B. subtilis

(DBTBS) or proteome databases (KEGG, UniProt and PDB) (Nicolas et al., 2012; Kanehisa et al.,

2016; Burley et al., 2017). SubtiWiki, a highly detailed database, has been developed to gather

together all information of B. subtilis, from specific databases of genes, messenger RNAs, protein

expression, protein-protein interactions to regulatory and metabolic pathways (Michna et al. 2014;

Zhu and Stülke 2018).

1.1.4 B. subtilis: a bacterium of industrial interest

In addition to its GRAS (Generally Recognized as Safe) status, B. subtilis and close relatives

are a source of products with industrial importance for the production of numerous proteins such as

amylases (used in the bread industry), proteases, cellulases (used in the detergent industry) or

antibiotics such as bacitracin and polymyxin (van Dijl and Hecker 2013; Westers, Westers, and Quax

2004; H. Cao et al. 2017). This capability is mainly due to two intrinsic characteristics of this group of

bacteria:

I. In their natural habitat, the soil, they have the ability to produce a wide range of hydrolytic

enzymes that allows them to breakdown soil-based macromolecules into vital nutrients (Pohl

and Harwood 2010b).

II. They are also able to export and secrete proteins directly into the extracellular medium at

concentrations as high as grams per litre, due in part to the single membrane system of Gram-

positive bacteria, in contrast to the double membrane system of Gram-negative bacteria.

The fact that B. subtilis is easy to grow at high cell densities and its natural ability to release

secretory proteins into the culture medium is of great industrial and commercial interest. It reduces

downstream processing costs, makes purification more straightforward and reduces the likelihood of

contamination with cytoplasmic proteins (van Dijl and Hecker 2013; H. Cao et al. 2017) .


18

Box2: B. subtilis

Commonly found in the soil, B.subtilis, is a rod shaped ubiquitous bacterium that

secretes numerous enzymes to the culture medium, enabling the bacterium to survive in a

continuously changing environment. It is considered as a GRAS organism, it can be easily

cultivated up to high cell densities and handled in the laboratory. Along with its high genetic

tractability and amenability to adopt different life-styles, B. subtilis has become the Gram-

positive model organism, fundamental for research studies, and a cell factory used for large-

scale production of industrial enzymes.


19

1.2 Protein translocation across cytoplasmic membrane in

bacteria

Membrane and exported proteins are crucial players for maintenance and survival of bacterial

organisms and their correct compartmentalization is a key step for molecular function (Figure 2).

After being synthesized by ribosomes in the bacteria cytoplasm, these proteins need to be inserted in

the cytoplasmic membrane, anchored to the membrane or released into the extracellular medium

(Green and Mecsas 2015; Schneewind and Missiakas 2012; Costa et al. 2015).

Figure 2. Protein export pathways in B. subtilis. Schematic representation of B. subtilis cell and its export

pathways. After the protein synthesis by the ribosomes, proteins undergo different destinations depending on the

presence (+SP) or absence (-SP) of an N-terminal signal peptide (SP) and its category. Proteins without a signal

peptide remain in the cytoplasm. Proteins that have a typical secretory signal peptide are exported via the Sec

pathway, but if the SP contains a twin-arginine motif (+RR) the protein is exported by the Tat pathway. These

proteins can be then secreted to the medium, retained at the membrane by a transmembrane segment (TM),

anchored to the extra-cytoplasmic side of the membrane by a lipid modification (+lipobox) or anchored to the

cell wall, the mature parts of these proteins contain cell wall-binding repeats (+CWB). Pseudopilins are exported

by the Com system. ABC transporters can export peptides too, as it is the case of pheromones and lantibiotics.

(H Tjalsma et al. 2004)

Exported proteins have important functions as, among others, hydrolysis of extracellular

polymers (proteins, nucleic acids, polysaccharides), synthesis of the cell wall and intercellular matrix,

such as biofilm, modification of the host cell, and sensing of the environmental conditions (Ivankov et

al. 2013). In the present chapter, I will describe all the export routes that membrane and secretory

proteins can undertake in bacteria, with greater emphasis on the Gram-positive B. subtilis.


20

1.2.1 The general protein secretion pathway: the Sec system

In bacteria, the Sec system is highly conserved and it represents the major route for protein

transportation through the hydrophobic membrane barrier. The Sec system allows the translocation of

proteins in an unfolded conformation, without loss of small molecules such as ions and valuable

metabolites. This system was originally identified in E. coli back in the early 90’s. By the use of

liposomes, a preprotein substrate (proOmpA) mixed with Sec purified proteins, respectively SecB,

SecA and SecY/E proteins, these early studies showed that these components form a binding cascade

from SecB to SecA to SecY/E and that they may be enough for the basic reactions involved in the

targeting and the translocation of a cargo protein (Brundage et al. 1990; Hartl et al. 1990). The Sec

translocase, in its minimal form, consists of a protein-conducting channel formed by the heterotrimeric

membrane protein complex SecYEG, and the essential ATPase SecA that acts as a molecular motor

mediating protein transport in this pathway (Brundage et al. 1990; Akimaru et al. 1991; Beckwith

2013).

In general, the main components of the Sec system are highly conserved, and homologous of

the SecYEG complex (Figure 3) have been found in the cytoplasmic membrane of archaea, the

chloroplast thylakoid membrane and the eukaryotic endoplasmic reticulum (Pohlschröder et al. 1997).

However, recent studies on the Gram-positive Sec system revealed that a large number of Gram-

positive bacteria possess accessory Sec components that are not found in other organisms. A second

SecA paralog, termed SecA2, and a second SecY paralog, termed SecY2, are present and although

being involved in protein translocation, in most cases, they are not essential (Prabudiansyah and

Driessen 2016). This accessory pathway is found in different Gram positive bacterial species as

Mycobacterium, Listeria, Clostridium difficile, Streptococcus, Staphylococcus and Bacillus. Bacillus

subtilis is devoid of accessory SecA2 protein, however this is present in some Bacillus species such as

Bacillus anthracis, Bacillus cereus, Bacillus thuringiensis, Bacillus smithi, and Bacillus methanolicus.

SecA2 promotes the export of a small subset of proteins and plays an important role in virulence

(Kurtz et al. 2006; Sullivan et al. 2012). SecY2 is predicted to form an accessory membrane channel

that is responsible for the export of specific proteins, which cannot be exported by the canonical

SecYEG (Prabudiansyah and Driessen 2016). These accessory SecA2/Y2 pathway appear to mediate

the transport of the surface protein GspB, the superoxide dismutase SodA and the catalase peroxidase

KatG (Bensing and Sullam 2002; Kurtz et al. 2006).


21

Figure 3. Schematic prediction of the B. subtilis Sec-dependent secretory protein translocase. A preprotein

substrate is pushed, in an unfolded state, through the heterotrimeric pore (SecYEG) by the SecA motor protein.

The SecDF–YrbF complex and the main type I signal peptidases (SipS and SipT) are involved in the substrate

translocation by helping on the translocation mechanism and cleaving the signal peptide, respectively. After

translocation, the preprotein is folded into its correct formation by Sec accessory proteins (folding factors).

Adapted from Harwood and Cranenburgh 2008.

Briefly, the translocation through the Sec pathway can be divided in to three main steps

(Figure 4):

I. Targeting: The secreted proteins are synthesised as precursors (pre-pro-protein) with a N-

terminal signal peptide (SP), which is recognized by cytoplasmic factors (Signal Recognition Particle-

SRP) or chaperones, such as SecB in E. coli and CsaA in B. subtilis (Collinson 2005; H Tjalsma et al.

2000). These factors are responsible for maintaining the substrate in an unfolded secretory state and

for targeting the pre-protein to the translocon formed by the SecYEG complex.

II. Translocation: The secreted or membrane proteins are pushed into the SecYEG channel. This

movement is driven by the translocon motor SecA, an ATPase that interacts with the substrate proteins

and binds to the SecYEG complex (Bauer et al. 2014). The repeated cycles of binding, hydrolysis of

ATP, conformational changes of the dimeric SecA leads to the translocation of the secretory protein

through the SecYEG complex (Robson et al. 2009).

III. Release and maturation: Subsequently, the signal peptide is cleaved resulting in the release of

the protein from the membrane. Finally the protein undergoes a maturation process carried out by

other accessory proteins, such as the foldase PrsA, or the Bdb thiol-disulfide oxido-reductases

(Bolhuis et al. 1999; van Wely et al. 2001; Driessen and Nouwen 2008).


22

Figure 4. Components involved in Sec-dependent protein export in B. subtilis. Secretory proteins are

ribosomally synthesized as precursor proteins with an N-terminal signal peptide (SP). Cytoplasmic chaperones,

such as SRP/FtsY, keep the precursors in a translocation-competent state and facilitate their targeting to the

translocase in the membrane, consisting of SecA, SecY, SecE, SecG, and SecDF. During or shortly after

translocation, the preprotein is cleaved by one of the type I signal peptidases (SipS-W) or lipid modified by the

diacylglyceryl-transferase (Lgt) and cleaved by the lipoprotein-specific signal peptidase (Lsp). SppA may be

involved in the degradation of cleaved signal peptides, whereas the folding of several secreted proteins depends

on the activities of PrsA, BdbBC, and/or SpoIIIJ/YqjG. HtrA, HtrB, and WprA are involved in the quality

control of secretory proteins. It should be noted that for reasons of simplicity, HtrAB are depicted in the cell

wall, although HtrA is detected in both the membrane and the medium. On passage through the cell wall, the

mature protein is released into the environment. Adapted fom Tjalsma, H., et al. (2004).

1.2.2 Substrate recognition

A crucial early event in the Sec secretion pathway is the identification of substrates that are

destined for secretion by cytoplasmic components. The first evidence of a substrate recognition system

for protein translocation dates back to the 70’s, when Milstein and his colleagues (1972) provided

insights into how secreted immunoglobulins were made in precursor form with an amino-terminal

hydrophobic sequence that was cleaved to the mature form of the protein, raising the signal sequence

hypothesis (Milstein et al. 1972). Few years later, Blobel and Dobberstein provided substantial and

persuasive evidences into how secreted proteins are recognized. They showed, through in vitro studies,


23

that the product synthesized from the mRNA for the light chain of lgG was larger by 4,000 g/mol than

the actual secreted light chain (Blobel and Dobberstein 1975).

The non-mature preproteins have a hydrophobic N-terminal extension, the signal sequence or

signal peptide (SP), that can be recognised by the targeting system (as SRP, SecB and CsaA)

responsible for directing the preproteins to the cytoplasmic membrane (Ivankov et al. 2013). The

signal sequences are typically between 15 up to 30 amino acids long and they are characterized by

three different domains:

i. N-domain: a 5- to 8- residue-long positively charged amino terminal that interacts with the

translocation machinery and the negatively charged phospholipids of the membrane;

ii. H-domain: a 8- to 12- residues-long hydrophobic core region decisive for the signal

peptide function. The presence of these hydrophobic residues promotes an α-helical

conformation and consequently a formation of a hairpin-like structure that might result in

the insertion of the signal peptide into the membrane;

iii. C-domain: a 5- to 7- residue long sequence containing a cleavage site for specific signal

peptidases (SPase) responsible for removing the signal peptide from the exported protein,

during or shortly after its translocation.

Although, the signal peptides have this structure in common there is a high variability of the

amino acids sequence Figure 5). In general, the signal peptides of Gram-positive bacteria are typically

longer and more hydrophobic than the SP of Gram-negative bacteria (Zanen et al. 2005; Harwood and

Cranenburgh 2008). Responsible for the targeting of membrane proteins and secretory proteins to the

correct secretion machinery. The SP can be classified according to the SPase cleavage sites and the

pathway by which the preproteins are predicted to be exported. The Sec-dependent SPs, which are the

most abundant type of SPs, contain a characteristic AXA motif that allows the targeting to the Sec

translocation machinery.


24

Figure 5. General features of the signal peptides and propeptides of Bacillus secretory proteins. The signal

peptides tripartite structure, N-terminal (N) containing lysine and/or arginine residues (indicated by +),

hydrophobic (H, indicated by a grey box) and cleavage (C) regions are schematically represented. The export

pathways via which the preproteins are exported and the SPases responsible for their cleavage are indicated. The

break between the boxes represent cleaved sites and the most frequently occurring first amino acid of the mature

protein (+1) is indicated. Tat-dependent signal peptides with twin arginine motif (SRRxFLK) and the Sec-

dependent signal peptides containing an AXA cleavage site, are cleaved by a type I signal peptidase (SPase).

Lipoprotein signal peptide cleaved by the type II SP. The pseudopilin-like signal peptides and bacteriocin and

pheromone signal peptides have specific SPases. The length of the signal peptides and their subdomains is drawn

to the same scale. (H Tjalsma et al. 2004)

The signal peptides present in B. subtilis can be classified into five distinct classes: twin-

arginine (SRRxFLK) signal peptides, secretory (Sec-type) signal peptides, lipoprotein signal peptides,

pseudopilin-like signal peptides, and bacteriocin and pheromone signal peptides (H Tjalsma et al.

2004). The Tat-dependent SPs have a twin arginine motif (RR/KR) specific that directs them to the

Tat secretion machinery (Natale, Bruser, and Driessen 2008). The difference between the SP of the


25

secretory proteins and lipoproteins is the presence of a conserved lipobox. This sequence contains a N-

terminal invariable cysteine residue that have to be diacylglycerol-modified before their attachment to

the outer surface of the membrane (Harwood and Cranenburgh 2008). In B. subtilis four proteins have

been identified with pseudopilins SPs (ComGC, ComGD, ComGE, and ComGG) which are cleaved

by the SPase ComC (Chung and Dubnau 1998). Finally, the fifth class of SPs refers to the ribosomally

synthesized pheromones and lantibiotics, which contain only a N- and C-domains and completely

devoid of a hydrophobic H-domain (Fig. 4). These types of proteins are exported and cleaved by ABC

transporters (Harold Tjalsma et al. 2000).

Since the SPs are an essential prerequisite for a successful targeting and secretion of secreted

proteins, it is a good starting point for the optimization of secretion of heterologous proteins. In

general, the signal peptides of Gram-positive bacteria are typically longer and more hydrophobic than

the SP of Gram-negative bacteria (Zanen et al. 2005; Harwood and Cranenburgh 2008). Zanen et al,

have showed that the hydrophobicity is critical for early signal peptide recognition. B. subtilis is not

able to secrete preproteins with signal peptides of low hydrophobicity but secretory proteins with

signal peptides with high hydrophobicity would be more efficiently translocated in this bacterium

(Zanen et al. 2005).

As an attempt to optimize the secretion of heterologous proteins, Brockmeier et al. (2006,)

created a library of all natural Sec type SPs from B. subtilis fused to reporter proteins that were then

tested with a screening approach, making use of two lipolytic enzymes- cutinase from Fusarium solani

pisi and a cytoplasmatic esterase of metagenomic origin. They observed that the best SP of one target

protein may not be the best or even work for other secreted proteins. As an example, the fusion of the

SP of AmyE to the cutinase did not inevitably result in an improvement of the secretion. As a

conclusion, the authors showed that the secretion efficiency for a certain target protein in B. subtilis is

determined by complex modifications and interactions between the signal peptide and the mature

protein (Brockmeier et al. 2006).

1.2.3 Intracellular chaperoning and targeting to the Sec translocase

The targeting of the secretory preproteins to the Sec pathway in bacteria is an important step

for a correct protein exportation. During this stage, preproteins interact with cytosolic chaperones to

undertake the correct route, to be maintained in the correct unfolded state and to ensure a driving force

that provides directionality to the secretion, because polypeptides can slide by Brownian motion in

either direction inside the channel (Rapoport, Li, and Park 2017). The protein targeting to the channel

can occur by two distinct routes, by a co-translational or post-translational route (Figure 6).


26

Figure 6. Schematic representation of the different modes of protein translocation across the membrane of

B. subtilis. (A) Different stages of cotranslational translocation. The polypeptide is moved directly from the

ribosome tunnel into the SecYEG channel. The ribosome is anchored to the membrane by the SRP-receptor

complex. (B) In the model for posttranslational translocation, the preprotein translocation is mediated by the

SecA ATPase.

1.2.3.1 Co-translational translocation route

In the co-translational path, the new emerging protein chain is simultaneously synthesized,

pushed directly from the ribosome exit tunnel through the translocon and exported or laterally inserted

into the lipid bilayer. This co-translational route is performed by the signal recognition particle (SRP),

which is a highly conserved ribonucleoprotein complex, in all three kingdoms of life, and which is

responsible for recognizing the hydrophobic signal sequence of the ribosome-nascent chain (RNC) as

it emerges from the ribosomes and targeting it to the cytosolic membrane (Koch, Moser, and Müller

2003). In general, SRP is composed of a small cytoplasmic RNA (scRNA) and signal sequence

binding subunit (Ménétret et al. 2005; Collinson, Corey, and Allen 2015). First, this protein complex

binds to a membrane receptor, forming a heterodimeric SRP-receptor complex, which helps the

targeting of the emerging nascent pre-protein from the ribosome to the membrane after the binding to

the signal peptide (Saraogi and Shan 2014). E. coli SRP comprises a scRNA domain and a single

essential protein, Ffh. After the recognition of the hydrophobic signal sequence of the emerging

protein, the SRP interacts with the SRP membrane receptor FtsY, which leads to the transfer of the

emerging protein to the Sec translocase for translocation or insertion into the membrane (Angelini,

Deitermann, and Koch 2005; S. Chen et al. 2008). Ffh and FtsY belongs to the SRP-GTPase family,

depending on GTP-binding, and both are essential for SRP-dependent protein secretion and cell

viability (S. Chen et al. 2008; Neher et al. 2008). B. subtilis has a co-translational route too. As the E.

coli SRP, B. subtilis SRP consists of a scRNA and a GTPase Ffh, however it additionally contains two

molecules of the Histone-like protein Hbsu (Nakamura et al. 1999). The SRP can be divided into two

functional domains: the S domain recognizes the N-terminal signal sequences as soon as the pre-

protein emerges from the ribosomal tunnel exit and the Alu domain imposes an elongation arrest by


27

blocking the elongation factor entry site, retarding translation (Kempf, Wild, and Sinning 2014;

Rodnina and Wintermeyer 2016). Therefore, by stopping the elongation, SRP prevents membrane

proteins from being prematurely released from the ribosome before the ribosome-nascent chain

complex (RNC) has correctly engaged with the translocation channel (Kempf, Wild, and Sinning

2014). In B. subtilis SRP, the HbsU protein binds to the SRP Alu-Like domain, being essential to this

process. In a study using β-lactamase as a reporter protein, the repression of hbs gene and depletion of

Hbsu leads to slow growth and defect in the translocation with a high accumulation of the preprotein

form of the β-lactamase and an induced scRNA upregulation (Yamazaki et al. 1999).

1.2.3.2 Post-translational translocation route

The post-translational translocation route involves the binding of specialized energy-

transducing factors, such as ATPases. Although in eukaryotes the translocation occurs across the

endoplasmic reticulum (ER) membrane and is realised by a larger complex formed by trimeric Sec61

complex associated with additional components, the post-translocational mechanism works as in

bacteria (Osborne, Rapoport, and van den Berg 2005). It needs a chaperone that acts as a molecular

clamp to grab the emerging polypeptide, directing to the pore and preventing its backsliding. This

mechanism occurs by cycles of hydrolyses of ATP, that consequently increase the affinity for

unfolded polypeptide (Collinson, Corey, and Allen 2015). The SecB protein is the secretory chaperone

in E. coli. This protein binds to the mature regions of a subset of preproteins maintaining them in a

translocation-competent state (unfolded and extended state) and to the carboxy terminus of SecA,

which suggests that SecB promotes protein translocation of the associated preproteins by its specific

binding to SecA. Therefore, the SP interacts post-translationally with SecA, the protein motor of the

pore SecYEG, that subsequently undergoes different conformational states and cycles of ATP and

ADP binding states promoting the translocation through the SecYEG pore by a “push and slide”

movement (Bauer et al. 2014). Then the SecD, SecF and YajC, associated membrane proteins,

prevents the slide back of the translocated protein (Corey et al. 2016).

SecB protein is absent from B. subtilis and all other Gram-positive bacteria, and many studies

have focused on identifying a functional homologue of this protein in this group of microorganisms.

To date these studies have failed to identify a convincing homologue. It might be important to note

that SecB is not essential for E. coli viability and it is only needed for the secretion of a specific subset

of proteins, many of which are outer membrane proteins (van Wely et al. 2001). The absence of outer

membrane proteins in Gram-positive bacteria might explain why a homologue of SecB has not yet

been found in this group of organisms. However, according to Fekkes et al. (1997) the B. subtilis

SecA protein contains a highly conserved binding domain that is present in SecB, which suggests that

SecA has a chaperone-like function (Fekkes, Van der Does, & Driessen, 1997). In 2012, Diao et al.

showed a significant improvement on the secretion of a maltose binding protein (MalE11) and an

alkaline phosphatase (PhoA) with the implementation of an artificial posttranslational protein targeting


28

pathway in B. subtilis by co-expressing SecB from E. coli together with a B. subtilis hybrid SecA

protein, in which the C-terminal 32 amino acids were replaced by the corresponding part of SecA from

E. coli (Diao et al. 2012). This data indicated that, in this route, SecA plays part of the role of SecB.

However, B. subtilis possibly has another homologous protein of SecB that contains other binding

domains that might serve as a docking site for the SecB analogue. The cytosolic CsaA protein is the

possible homologue of SecB in B. subtilis. It was demonstrated, that, a CsaA has a chaperone-like

activity both in vitro and in vivo, with affinity for SecA and preproteins, as well as a role in the protein

secretion, as the suppression of the csaA gene leads to a decrease of the secretion efficiency in B.

subtilis (Müller et al. 2000). Furthermore, it was shown that the intracellular and general chaperones

GroES and GroEL or DnaK, DnaJ and GrpE can work on pre-proteins and maintain them in a

translocation-competent form. By inactivating the gene encoding HrcA, a negative regulator of the

groE and dnaK operons, the authors bserved that the synthesis of all these chaperones. This increases

the yields of the single-chain antibody (SCA) fragment and a reduced inclusion body formation (S. C.

Wu et al. 1998).

1.2.3.3 SecA and the Sec translocase

In B. subtilis, the membrane bound SecYEG complex forms the translocon (De Keyzer, Van

Der Does, and Driessen 2003). As the name implies, this complex includes the SecY, SecE, and SecG

proteins. SecY is the largest component of the translocase and is essential for translocation and

viability. The secY gene is located within a ribosomal operon, the spc operon. SecY is predicted to

span the cytoplasmic membrane 10 times and is found in randomly dispersed foci that dynamically

assemble and disassemble in the membrane (Dajkovic et al. 2016). SecE is a small integral membrane

protein that is essential for translocation and viability (Jeong, Yoshikawa, and Takahashi 1993). In E.

coli, the SecE three transmembrane domains (TM), surround the back of SecY and appear to stabilize

SecY by preventing it from being degraded by FtsH, an ATP-dependent protease present in the

cytoplasmic membrane (Lycklama A Nijeholt et al. 2013). As mentioned before, SecE is part of the B.

subtilis translocon, however experimental evidences do not exist that it has the same function than in

E. coli. It is assumed that this small protein has the same function because of its homology to E. coli

SecE. SecG is the third component of this integral membrane complex. It is the only component that is

not essential for the cell viability and the protein translocation. In B. subtilis, the deletion of this gene

causes cold sensitivity when secretory stress is imposed (van Wely et al. 1999). These observations

suggest that SecG only contributes to the efficiency of the translocation reaction. During the

translocation cycle, the SecYEG translocon assume different conformational stages (Zheng et al.

2016).

SecA is an ATPase located on the cis side of the membrane which is responsible for both the

preprotein targeting and the coupling of the energy required for translocation. It is the motor

component responsible for pushing the preprotein through the SecYEG pore, as illustrated on


29

Figure 7 (Bauer et al. 2014). Therefore, SecA is considered as part of the core of the

translocation channel as the motor protein of the SecYEG translocon interacts with both proteins

involved in the translocation, the secretory preprotein and the membrane proteins that compose the

translocon (Bauer et al. 2014; Tanaka et al. 2015).

Figure 7. Scheme of SecA-mediated protein translocation. (A) SecYEG closed channel; (B) Formation of the

SecA–SecY complex; (C and D) At this stage the channel is active and the translocation starts; (E) The final

stage where the signal peptide is cleaved and the protein is released. The translocating polypeptide is shown in

green and the signal peptide as a green cylinder. The lateral gate of the channel is shown as a broken line on the

left. (Alguel et al. 2016)

Kakeshita et al. 2010 showed that the SecA C-terminal region is not conserved in eubacteria

and by deleting 61 amino acids of this region led to significant enhanced production of heterologous

proteins in B. subtilis. The membrane translocase complex itself is of great interest for the

optimisation of production strains. In an attempt to solve the problem of jamming of the membrane

caused by the overexpression of secretory proteins through the increase of the number of translocon,

Mulder et al (2013), constructed an artificial SecYEG operon under the control of an inducible

promoter. They demonstrated that the yield of α-amylase could be improved by increasing the

production of the SecYEG proteins (Mulder, Bandola, and Schumann 2013). To investigate the

bottleneck of the Sec pathway in B. subtilis Chen et al. (2015) developed a combinatorial approach to

evaluate the effect of a systematic overexpression of 23 genes or operons involved or related to this

route for the production of AmyL and AmyS. Contrary to Mulder et al. (2013) results, this new

approach showed that the overexpression of the translocase genes did not significantly increase the

production of heterologous proteins, but the overexpression of post-translocation molecular chaperone

(PrsA) leads to an optimization of AmyL and AmyS secretion (J. Chen et al. 2015). Furthermore, in

2016, Dajkovic and colleagues revealed by internal reflection microscopy (TIRFM) that SecA and

SecY form foci throughout the cytoplasmic membrane. These foci were in constant flux with freely

diffusing SecA and SecY molecules, concluding, this way that the secretion system is remarkably

dynamic and is in constant reorganization (Dajkovic et al. 2016).


30

1.2.3.4 Protein release and maturation

The final step of the Sec translocation consists in the cleavage of the signal peptide (SP) by

Signal peptidases (SPases), resulting in the release of the protein from the membrane to the media or

lipid modified by the diacylglyceryl-transferase and cleaved by the lipoprotein-specific signal

peptidase. SPases are membrane proteins responsible for the cleavage of the signal peptide and they

are divided in Type I, Type II and TypeIV (Figure 8).

In bacteria, SPaseI cleaves the majority of SP of the secretory pro-proteins, mainly proteins

released to the medium or cell wall proteins. Lep is the only type I signal peptidase that E. coli has,

while B. subtilis has five: SipS, SipT, SipU, SipV, and SipW. SipS and SipT are the main signal

peptidases and their deletion leads to cell death (Harold Tjalsma, Van Den Dolder, et al. 1999).

SPasesI are general signal peptidases, catalysing the release of exported proteins through the Sec

pathway and, also, by the Tat pathway, which facilitates the export of fully folded proteins (Lüke et al.

2009). These peptidases, that have a large C-terminal domain characterized by two transmembrane

helixes and an active site extended to the extra-cytoplasmic membrane surface, are unusual serine

protease, as the active site exhibits a Ser-Lys dyad configuration instead of the canonical Ser-His-Asp

triad architecture (Dalbey, Wang, and van Dijl 2012).

The SPaseII, the lipoprotein signal peptidase, has a crucial role in recognizing and processing

the lipid-modified bacterial membrane proteins, which have to be diacylglycerol-modified at an N-

terminal cysteine residue prior to their attachment to the outer surface of the membrane (Pohl and

Harwood 2010). In B. subtilis, Lgt is the diacylglyceryl-transferase responsible for this lipid

modification, prior to the SP cleavage by the SPaseII Lsp, contrary to E. coli this protein is not

essential for cell viability (Harold Tjalsma, Zanen, et al. 1999). This peptidase is an aspartic acid

protease, whose catalytic residues are fixed just beneath the extra-cytoplasmic membrane surface.

The SPaseIV, the prepilin signal peptidase, are responsible for the processing of the precursors

of type IV pilins and related pseudopilins involved in the secretion of proteins across the outer

membranes of Gram-negative bacteria (type II secretion) or in DNA uptake by Gram-positive bacteria.

In contrast to the other types of peptidases, this type has a specific recognition sequence, Gly-Phe-Thr-

Leu-Ile-Glu, where cleavage occurs between the Gly and Phe residues, and is also responsible for N-

methylation of the phenylalanine at position +1 relative to the cleavage site. Similar to SPaseII, these

peptidases are intramembrane-cleaving aspartic acid proteases, characterize by eight transmembrane

helixes with the active site positioned on the cytoplasmic side close to the membrane, within short

cytoplasmic loops (Strom, Bergman, and Lory 1993).


31

Figure 8. Signal peptide cleavage of precursor proteins by signal peptidases. Signal peptidase I (SPaseI)

employs a Ser-Lys catalytic dyad for signal peptide cleavage from secretory precursor proteins at the

extracytoplasmic surface of the membrane. The Protein Data Bank (PDB) structure of the catalytic domain

(accession number 1T7D) and the program JMol were used to generate the three-dimensional (3D) structure

image of SPI. Signal peptidase II (SPII) is an aspartic acid protease that cleaves signal peptides from bacterial

lipoprotein precursors just beneath the extracytoplasmic membrane surface. Signal peptidase IV (SPIV) is an

aspartic protease that cleaves signal peptides from prepilins and pseudopilins at the cytoplasmic surface of

bacterial membranes. Transmembrane helices of signal peptidases are depicted as blue barrels, and substrate

helices are depicted as red barrels. A zoomed-in view of the active site residues of SPI is shown. The locations of

the N and C termini of the signal peptidases and their substrates are indicated. Dalbey, Wang, and van Dijl 2012.

Beyond the cleavage of the signal peptide and subsequent release of the mature protein, there

can be other limitation step for the optimization of the secretion, which implies an optimization of the

rate of the signal peptides processing by the signal peptidases. Therefore, Tjalsm et al (2000)

described different signal peptidases present in B. subtillis which have different affinities for different

substrates, suggesting that the overexpression of the right combination of signal peptides will lead to

an increase of the protein of interest (Harold Tjalsma et al. 1997; H Tjalsma et al. 2000). The rate of

signal peptide processing by signal peptidases has also been considered as a possible limiting factor

for protein production. Vitikainen et al. (2001) showed that the proportion of mature AmyQ in SipT-

overproducing cells was higher than in the wild type (Vitikainen et al. 2001).

After this cleavage reaction the released SP need to be recycled, otherwise these peptides

could have an inhibitory action to protein translocation. SppA and RasP have been described as

possible signal peptide peptidases responsible for this degradation in B. subtilis (Bolhuis et al. 1999;

Neef et al. 2017).

Gram-positive bacteria are known by their characteristic cell envelope architecture that

facilitates the release of secretory proteins directly into the culture medium. Therefore, it is expected

that this architecture would be compatible with the protein export process. However, a closer look at

the cell wall reveals that it offers a very challenging environment for proteins to fold after emerging

from the translocase. Unfolded proteins that emerge from the translocase must be able to fold into


32

their native configuration, a crucial step not only for their functionality but also for their stability. The

folding process should be quick without forming intra- or inter-molecular interactions that could block

the translocation machinery and/or block cell wall synthesis, either of which would compromise cell

viability. B. subtilis contains several post-translocational and quality control proteases in the

membrane and cell wall, as well as extracellular proteases that rapidly target heterologous proteins or

misfolded native proteins with exposed protease recognition sites. In B. subtilis, key roles are played

by PrsA, the folding catalyst, various thiol-disulphide oxidoreductases (BdbA, BdbB, BdbC and

BdbD), cleaning proteases such as the membrane-associated HtrA and HtrB proteases and

extracellular proteases as WprA.

PrsA is a peptidyl-prolyl cis-trans isomerase (PPIase), an essential membrane anchored

lipoprotein, bound to the outer surface of the cytoplasmic membrane, that assists proteins with cis-

prolyl residues to fold (Kontinen, Saris, and Sarvas 1991; Kontinen and Sarvas 1993). Its deletion

results in altered cellular morphology and cell death given that this protein is required for the folding

of one or more proteins involved in cell wall synthesis (Kontinen and Sarvas 1993). Moreover, several

studies have revealed that there is a linear relationship between the amount of PrsA and the yield of

certain secretory proteins (notably amylases), making PrsA a popular target when engineering host

strains dedicated to the production of native, as well as heterologous secretory protein (Vitikainen et

al. 2005).

Due to the oxidized environment into which secretory proteins are translocated, wich favours

disulfide bond formation, B. subtilis expresses four thiol-disulphide oxidoreductase/isomerase

enzymes (BdbA, BdbB, BdbC and BdbD) involved in the disulphide bond formation (Sarvas et al.

2004). However, secretory proteins from B. subtilis and other Gram-positive bacteria generally lack

disulphide bonds. Nevertheless, these enzymes have an important role in the production of

heterologous proteins that require such bonds.

HtrA and HtrB are serine proteases, belonging to the quality control protease family,

combating the extracellular folding stress, and preventing potentially fatal obstruction of the secretory

translocase and cell wall synthesis (Darmon et al. 2002; Antelmann et al. 2003). It was recently

suggested that, in addition to their already described roles as quality control proteases, HtrA and HtrB

might also function to release lipoproteins from their membrane anchorage, to degrade membrane

proteins and also to facilitate the folding of native secretory proteins (Krishnappa et al. 2013).

WprA is one of the eight extracellular proteases encoded in B. subtilis (AprE, Bpr, Epr, Mpr,

NprB, NprE, Vpr), which is also localized on the cell wall (Krishnappa et al. 2014). It is a serine

protease involved in the degradation of peptides for recycling and nutrient provision and the

proteolytic processing of other proteins (Babe and Schmidt 1998). Although it was believed that,

correctly folded Sec-secreted proteins would be highly resistant to the extra-cytoplasmic proteases,

Krishnappa et al. (2014) have shown that many of these proteins are in fact degraded. They showed


33

that the deletion of WprA leads to an increase in the amount of PrsA secreted into the medium,

demonstrating that WprA has a role in the degradation of PrsA (Krishnappa et al. 2014).

1.2.4 Twin-Arginine Transport (Tat) Pathway

In contrast to other transport systems, the twin-arginine translocase (Tat) is an unusual

protein-targeting export pathway, found in all the kingdoms of life, that uniquely facilitates the

translocation of fully folded and cofactor-attached proteins across the cytoplasmic membrane (Sargent

2007; B C Berks 2015). The active translocation is driven only by the transmembrane proton motive

force (PMF) (Anné, Economou, and Bernaerts 2016; Patel, Smith, and Robinson 2014). This system

was identified for the first time in the thylakoids of plant chloroplasts in 1997, it was shown that a

metallo-enzyme was translocated across the thylakoid membrane in a folded conformation through a

∆pH-dependent pathway (Settles and Martienssen 1998). This pathway has been evolutionarily

conserved in plant chloroplasts where it mediates protein import across the thylakoid membrane and is

essential for the formation of a functional photosynthetic apparatus (Alcock et al. 2016). Later in

1998, Weiner and his colleagues proved the existence of this pathway in prokaryotes. They identified,

in E. coli, a Sec-independent pathway which is required for the correct localization of periplasmic

redox proteins and for the targeting of DMSO reductase to the membrane (Weiner et al. 1998).

Consequently, Tat pathway plays a key role in a range of different cellular processes, because it

mediate the transport, in its majority, of proteins that bind cofactors in the cytoplasm and, thus, fold

Box3: Sec Pathway

The Sec pathway is highly conserved in bacteria and represents the main route for

protein transportation. It mediates the translocation of secretory and membrane proteins in

an unfolded conformation. This secretion process can be divided into three steps: targeting,

translocation and release. The secreted proteins are synthesised as precursors (pre-pro-

protein) with a N-terminal signal peptide (SP), which is recognized by cytoplasmic factors

(Signal Recognition Particle- SRP) or chaperones (CsaA) involved in targeting of the pre-

protein to the translocon SecYEG. The translocation of these proteins across the channel is

driven by the translocon motor SecA, an ATPase that interacts with the substrate proteins

and binds to the SecYEG complex. The repeated cycles of binding, hydrolysis of ATP,

conformational changes of a dimeric SecA leads to the translocation of the secretory protein

through the SecYEG complex. Subsequently, the signal peptide is cleaved resulting in the

protein release from the membrane. Finally, the protein undergoes a maturation process

carried out by accessory proteins, such as the foldase PrsA, or the Bdb thiol-disulfide oxido-

reductases (Driessen and Nouwen 2008; Kang et al. 2014; H Tjalsma et al. 2004).


34

before transport. Such co-factored proteins have essential roles in photosynthetic and respiratory

energy metabolism, iron and phosphate nutrition, resistance to heavy metals and antimicrobial

peptides, cell separation, symbiotic nitrogen fixation and membrane biosynthesis (Ben C. Berks,

Palmer, and Sargent 2003; De Buck, Lammertyn, and Anné 2008).

Similar to the Sec pathway, Tat substrates are targeted to the Tat apparatus by an N-terminal

signal peptide. Although Tat and Sec signal peptides share a similar overall architecture (Figure 9),

Tat signal peptides specifically exhibit a larger hydrophobic region, a general low hydrophobicity and

bear a conserved S/TRR-x-FLK (with x representing a polar aminoacid) eponymous motif that

contains the characteristic pair of adjacent arginine residues (twin arginines)(Alcock et al. 2016;

Goosens et al. 2015). A fact that gives to this pathway its name. The twin arginine residues are critical

for the Tat translocation in chloroplasts, but may not be an absolute prerequisite in bacteria, as the rate

of translocation is affected by mutating a single arginine, whereas mutation of both completely

abolishes the transport.

Figure 9. Tat pathway components. (A) Schematic representation of the Sec signal peptide and Tat signal

peptide. The different regions of the respective signal peptides (N-terminal (N), hydrophobic (H) and cleavage

(C) domains), mature protein parts, and conserved SP recognition sites are indicated. The black arrow indicate

the SPase cleavage site. (B) Illustration of the basic components of the Tat apparatus. The large transmembrane

protein TatC is shown in purple and the TatA-like proteins TatA, TatB and TatE are shown in pink, green and

orange respectively. Adapted from (Dalbey, Wang, and van Dijl 2012; Goosens, Monteferrante, and Van Dijl

2014).

In general, two types of integral membrane proteins compose the Tat translocon

(Monteferrante et al. 2013; Goosens et al. 2015; Alcock et al. 2016):

TatC: a highly hydrophobic and relatively large membrane protein with six transmembrane

helices, whose N- and C-termini faces the cytoplasm;

TatA-like proteins (TatA, TatB or TatE): smaller proteins with a single hydrophobic

transmembrane domain (TMD) at the N-terminus immediately followed by an amphipathic helix


35

(APH) and a C-terminal tail densely charged. The TatA-like components resulted from gene

duplication and they have evolved towards different functions, namely substrate binding (TatB)

and pore formation (TatA). E. coli possesses a third TatA paralog, TatE, which is functionally

equivalent to TatA but is present at a much lower concentration in the cell and is not essential for

Tat transport.

The Tat transport requires (i) a docking complex composed by TatB and TatC, which together

form a multivalent receptor complex that binds Tat substrates (Fröbel et al. 2012); and (ii) a pore

complex, which results from the assembly of multiple protomers of TatA (Gohlke et al. 2005).

Translocation is initiated when a substrate protein with the correct signal peptide is recognized and

bound to the docking complex located in the membrane (Figure 10). As soon as it binds, the TatC

component of the docking complex inserts the substrate proteins into the membrane (Blümmel et al.

2017). This binding event triggers the proton motor force (PMF) and, consequently, the recruitment of

pore-forming TatA protomers from a pool in the membrane to form the active TatABC-containing

translocation site (Sargent 2007; Alcock et al. 2016; Wojnowska et al. 2018). Upon completion of

transport, the substrate is released from the translocation site and the signal peptide is cleaved by

signal peptidase.

Figure 10. Gram-positive bacterial Tat pathway. Schematic representation of the envelope structure Gram-

positive bacteria and its Tat translocation machinery. Tat substrates can be targeted to the membrane, the cell

wall or the growth medium. Adapted from Goosens 2014.

Crosslinking and fluorescent microscopy experiments, showed the dynamic oligomerization of

TatA. Particularly, it was possible to visualize directly the dynamics in E. coli through the fusion of

TatA with a green fluorescent protein (GFP). When substrates are scarce, TatA is localized in the

membrane surrounding the cells, while it is found in large oligomers when the substrate is present in

high concentrations (Leake et al. 2008; Rose et al. 2013).

The Tat machinery transports folded proteins of widely varying size across the membrane,

these substrates may vary from 20 Å up to 70 Å of diameter comparing to an unfolded polypeptide

chain with 12 A° diameter transported by the Sec pathway (Gohlke et al. 2005; B C Berks 2015). The


36

variation of the TatA channel diameter according to large or small substrates, ensuring a tight seal

substrate transportation across the membrane without an uncontrolled ion leakage, it is a challenge

faced by the Tat machinery. The membrane-weakening and pulling mechanism hypothesis explains

how the Tat system manage this process. The model suggests that the N-terminal of the TatA

oligomers, that form the Tat translocon site, are capable of destabilize the membrane by weakening or

even rupturing the membrane upon substrate-binding, allowing the TatC component to pull the

substrate through the destabilized membrane (Frobel et al. 2012).

In 2013, Rodriguez and colleagues provided the first experimental evidence supporting this

model, schematically presented on Figure 11. They showed by NMR that E. coli TatA forms a ring-

like complex in the presence of the detergent C12E9, where the transmembrane helices form a ring

inside the membrane from which the APH extends allowing the formation of complexes with variable

subunit numbers. They also showed by molecular dynamics simulation that the periplasm side of the

lipid bilayer surrounding the TatA complex was destabilized because of the unusual short TMD

(Rodriguez et al. 2013).

Later in 2018, Hou and colleagues demonstrated in vivo that in the presence of a Tat substrate

the N-terminus TMD of TatA can destabilize the membrane. However in the absence of substrate, the

full length TatA counterbalances such weakening effects (Hou et al. 2018). Therefore, they proposed a

two-state model: (i) the activated state - in which the binding of the substrate to the TatABC complex

promotes the position switch of the TatA APH, which locally weakens the membrane, allowing the

substrate translocation across the membrane; (ii) the resting state- in the absence of the substrate, the

N-terminal membrane part of the APH compensates for distorting effects of the short TatA membrane

TMD anchor (Hou et al. 2018).

Figure 11. Model of the regulation of TatA pore formation by TatBC and substrate. (A) The inactive

monomeric form of TatA accommodates the depth of the membrane by tilting the short TMH, increasing the

angle between the TMH and the APH, and deeply inserting the APH into the membrane. (B) Interactions with

TatBC/substrate complex force an orientation of the TatA TMH roughly parallel to the membrane normally

favoring TatA polymerization. The N terminus is pulled into the membrane thinning and distorting the bilayer

in the pore and sensitizing the membrane to disruption. The TatA APH serves as a platform for the substrate that

is tethered to the TatBC complex by a signal peptide. In the drawing the TatA oligomeric ring is represented by

two opposing subunits. (C) Pulling of substrate protein into the membrane forces TatA subunits apart and

ruptures the membrane (Rodriguez et al. 2013).

In the case of Gram-positive bacteria as the actinomycetes (including the Streptomycetes,

Mycobacterium and Corynebacterium), the Tat system is similar to the Gram-negative bacteria and is


37

composed by TatA, TatB and TatC components (Anné et al. 2014). In contrast, Staphylococci and

Bacillus have a smaller core Tat pathway, mainly composed by a single pair of TatA and TatC and

devoid of TatB. B. subtilis have genes that encode for two TatC (TatCd and TatCy) and three TatA

(TatAd, TatAy and TatAc). These Tat components can assemble into two distinct Tat translocases that

operate in parallel, working independently of each other and translocating specific substrates. Both

systems appear to be localized at the cell poles and division sites (Monteferrante et al. 2013). The first

of the two systems, TatAy-TatCy (TatAyCy), was expressed constitutively over many tested

conditions and exports several substrates, including YwbN, an iron-dependent DyP-peroxidase, the

Rieske iron-sulfur protein QcrA and the alkaline phosphatase YkuE. In contrast, the second system,

TatAd-TatCd (TatAdCd), whose the only known substrate is the phosphodiesterase (PhoD), is only

expressed in conditions of phosphate limitation (Goosens et al. 2015; Patel, Smith, and Robinson

2014; Jongbloed, van der Ploeg, and van Dijl 2006).

In bacteria the Tat and Sec pathways work in parallel. Even if their systems have specific

components, they share associated proteins, allowing a crosstalk between the cellular systems

(Goosens, Monteferrante, and Van Dijl 2014). This have been documented in E. coli, the production of

heterologous membrane bound Rieske iron–sulfur protein, depends on the secretion of both pathways,

as a portion of this protein was shown to use a Sec–YidC-dependent membrane insertion mechanism

while the remaining portion was translocated in a Tat-dependent manner (Keller et al. 2012). In B.

subtilis, the TatAy–TatCy membrane complex interacts with the cell wall and quality control protease

WprA. Through yeast two-hybrid experiments, it was shown that WprA resulted in a decrease of the

amount of the TatAy–TatCy -translocated EfeB and YkuE proteins, suggesting a functional WprA–

TatAy–TatCy–EfeB/ YkuE interaction (Monteferrante et al. 2013).

Box4: Tat pathway

The twin-arginine translocase (Tat) is a unusual protein-targeting export pathway,

found in all the kingdoms of life, that uniquely facilitates the translocation of fully folded and

cofactor-attached proteins across the cytoplasmic membrane (Sargent 2007; B C Berks 2015).

An event, that remarkably has no requirement for ATP as energy source, the active

translocation is driven only by the transmembrane proton motive force (PMF) (Anné,

Economou, and Bernaerts 2016; Patel, Smith, and Robinson 2014). This pathway have this

name due to its signal peptides distinguish themselves by bearing a conserved S/TRR-x-FLK

(with x representing a polar aminoacid) eponymous motif that contains the characteristic pair

of adjacent arginine residues (twin arginines)(Alcock et al. 2016; Goosens et al. 2015). The

variation of the Tat machinery pore is remarkable, since it is able to transport folded proteins

of widely varying size across the membrane, from 20 Å up to 70 Å of diameter compared to

unfolded polypeptide chains with 12 A° diameter transported by the Sec pathway (Gohlke et

al. 2005; B C Berks 2015).


38

1.2.5 Flagella Export Apparatus (FEA)

Bacteria are motile microorganisms. To survive they need to swim or swarm depending on

their environment. The bacterial motor that allows this movement is called the flagellum, a rotating

semi-rigid helical filament. The flagellum was observed for the first time in 1681, when Antony van

Leeuwenhoek, the father of protozoology and bacteriology, observed his own faeces (Dobell 1932). In

one of his letters describing this observations, he clearly described a flagellated microscopic parasite

named two centuries later as Giardia intestinalis, and wrote that this “little animal” has “sundry little

paws” capable of strongly stirring the medium, a clear observation of flagella (Dobell 1932).

The bacterial flagellum is a highly efficient molecular nanomachine with approximately 120-

140 nm of diameter and about 10 to 20 µm long, composed by three main structures : the basal body,

the hook and the filament, which are made of about 25 different proteins (Erhardt, Namba, and Hughes

2010). The basal body, a membrane-anchored structure that works as a structural anchor, a reversible

rotary motor and flagellar protein export system (Minamino, Imada, and Namba 2008). The hook is a

curved, thicker and flexible cylinder that joins the basal body to the filament, contributing to the

synchronized rotation of the several flagella on the cell (Depamphilis and Adler 1971). The filament is

a helical propeller composed by thousands of copies of the protein monomer flagelin (F. Wang et al.

2017). All the components of these protein complexes are exported by the flagellum protein export

apparatus, using ATP and PMF as the energy source (Forster and Marquis 2012; Diepold and

Armitage 2015). The flagellum is a great example of how membrane protein super-complexes are of

importance for cell integrity.

Due to cell envelope differences, Gram-positive and Gram-negative flagellum are structurally

different. As the basal body is a transmembrane structure that attaches the filament and, the hook to

the cell, it is the structure that has more differences between the two types of bacteria. While the

Gram-negative bacteria have a longer basal body composed of four rings – the cytoplasmic disk (C-

ring), the membrane disk (M-ring), the periplasm disk (P-ring) and the outer membrane disk (L-ring);

the Gram-positive bacteria have only two basal body rings, a M-ring and a P-ring, as shown on Figure

12 (Erhardt, Namba, and Hughes 2010; Mukherjee and Kearns 2014). As I am reviewing the B.

subtilis secretion systems, I will describe in detail the flagellar protein export system.

(Berg 2003)


39

Figure 12. Bacterial flagella structure. Schematic representation of the flagellar motor of Gram-negative (A)

and Gram-positive (B) bacteria. Electron micrographic reconstruction of the flagellar basal body side view (C).

All the components that compose the flagellar motor are specified on the Figure 13. Adapted from Berg 2003.

As described above, the flagellar membrane complex, named basal body, works as a rotary

motor and flagellar protein export system, the two key elements for the existence and well-functioning

of the flagellum. The B. subtilis flagellar motor comprises an inner membrane FliF ring that sits on top

of a cytoplasmic gear-like rotor complex composed by FliG, a C-ring protein complex FliM and FliY

responsible for controlling the flagellar direction, and the stator transmembrane complex composed by

MotA and MotB proteins (Depamphilis and Adler 1971; Minamino, Imada, and Namba 2008). The

stator complex allows the flow of protons across the membrane. This flux leads to a conformational

change of MotA, which promotes the electrostatic interaction with the C-terminus of FliG, generating

the torque for flagellar rotation (Blair 2003; Berg 2003). Through FliF, the torque movement is

transferred to the rod complex, a conduction duct that drives the rotation from the basal body to the

hook and filament (Gabel and Berg 2003; Mukherjee and Kearns 2014). B. subtilis encodes five

homologs of rod proteins - FlgB, FlgC, FlhO, FlhP and FliE. The four proteins, FlgB, FlgC, FlhO and

FlhP, were co-purified with the B. subtilis hook–basal body, results that support their role in the rod-

shape structure (Depamphilis and Adler 1971; Kubori et al. 1997). The study of mutants depleted for


40

FlhO and FlhP showed that these two proteins are part of the upper part of the rod structure, close to

the hook assembly, since their deletion leads to the secretion of the hook subunits, to the media,

abolishing the hook formation (Courtney, Cozy, and Kearns 2012). In contrast, the deletion of fliE

does not abolish the hook formation, but reduces rod and hook proteins secretion. Therefore, FliE is

not only a structural component of the rod, but a regulatory component necessary for full and correct

secretion of the flagellar components (Burrage, Vanderpool, and Kearns 2018).

The flagellar protein secretion system translocates the flagellar components, across the

membrane, in an unfolded conformation, to the channel of the growing flagellum (a structure with 20

Å of diameter) to maintain the proteins unfolded while they move to the flagellum tip up to 10-20 µm

long (Evans et al. 2013). This translocation process requires cellular energy, using ATP and PMF as

the energy source.

Figure 13. B. subtilis flagellum. Cartoon depicting the putative structure of the B. subtilis flagellum based on

empirical data and similarity to S. enterica. Peptidoglycan is indicated in light gray. Membrane is indicated in

dark gray. Flagellar components are colored and labeled. Micrographs are superresolution fluorescence

micrographs of the indicated structures in wild type cells: filament (maleimide-stained HagT209C), hook

(maleimide-strained FlgET123C) and basal body (FliMGFP fusion protein). Adapted from Mukherjee and

Kearns 2014.

Despite their differences, the flagellar secretion apparatus and the type III secretion system

(T3SS) of pathogenic bacteria, the injectisome, are remarkably conserved. These two systems are

structurally similar and their secretion mechanism are analogous, and most part of the membrane and

ATPase complex components share sequence homology (Carpenter, Zuberi, and Ordal 1993; Diepold

and Armitage 2015; Deng et al. 2017).

The secretion machinery comprises a membrane gate complex, inserted into the FliF M-ring

and composed by FlhA, FlhB, FliP, FliQ, and FliR, five higly conserved transmembrane proteins, and

a cytoplasmic ATPase complex formed by FliH, FliI, and FliJ (Erhardt, Namba, and Hughes 2010).

The transmembrane protein FliO, part of this apparatus too, is responsible for an efficient assembly of

the export gate complex. FliP and FliR form the core structure of the gate complex, a starting point for


41

the assembly of the remaining components (Figure 14) (Fukumura et al. 2017). Recently, by

performing purification experiments, it was shown that salmonella FliP forms a homohexamer

dependent on the FliO complex help, their interactions are required for a FliP correct assembly

(Fukumura et al. 2017).

Figure 14. Model for the assembly process of the Gram-negative flagellar type III export apparatus. The

export apparatus is composed of a transmembrane export gate complex made of FlhA, FlhB, FliP, FliQ, and FliR

and a cytoplasmic ATPase ring complex consisting of FliH, FliI, and FliJ. The FliP dimers form a homohexamer

with the help of the FliO complex, followed by the assembly of FliQ, FliR, and FlhB and finally of FlhA during

MS ring formation in the cytoplasmic membrane. Then, the FliM/FliN complex binds to FliG to form the C ring

on the cytoplasmic face of the MS ring. Finally, the FliH/FliI/FliJ ATPase ring complex is formed at the flagellar

base through interactions of FliH with FlhA and FliN (FliY in B. subtilis), allowing export of substrates though

the central cavity of the FliP6 ring complex (Fukumura et al. 2017).

FlhA and FlhB are responsible for coordinating the flagellar protein export, through their C-

terminal cytoplasmic domains, and form a docking complex platform for the substrates, ATPase

complex (FliH, FliI and FliJ) and chaperones associated to this system (Fukumura et al. 2017). The

Salmonella and E. coli FlhA forms a homononamer. In E. coli, it was shown that this protein plays

also a key role in the energy transduction mechanism, acting as an ion channel to conduct H+ and

Na+. FlhA works along with the cytoplasmic ATPase complex or as a genetic backup that

compensates for the loss or inactivation of the ATPase complex (Minamino et al. 2016).


42

Figure 15. An entropic chain mechanism for flagellum growth outside the cell. Subunit crystallization

beneath the cap foldase provides a strong anchor (force to break anchor, FA). Sequential subunits are linked

(force to break link, FL) head-to-tail in a chain by juxtaposed terminal helices forming parallel coiled-coils.

Subunits docked at the export ATPase are unfolded and docked by the export machinery. The N-terminal helix

of the docked subunit is then captured (force to break docking, FM) into the subunit chain by the free C-terminal

helix of an existing subunit in the flagellar channel. Adapted from Evans et al. 2013.

During the flagellum growth, the exported substrates remain unfolded while they move

through the flagellum channel, from the bottom to the end, it is other example of the great importance

of protein-protein interactions. Evans et al. (2013) demonstrated that this mechanism uses the entropic

force of each unfolded substrate and that these substrates gather together thus creating a multi-subunit

chain through the flagellum channel by the isolation of the growing filaments (Evans et al. 2013).

Through these experiments, they also revealed that, each subunit is linked head-to-tail to the other via

an interaction between the juxtaposed amino N-terminal and carboxyl C-terminal helices, as it is

possible to see on Figure 15 (Evans et al. 2013). The first subunit on the filament bottom is docked to

the export machinery and its C-terminal is free to be captured by a free unfolded subunit, through

sequential rounds of linkage. Hence, a multi-subunit chain is created until the flagellum tip, where it is

predicted that the N-terminal of each unfolded subunit crystallizes (Evans et al. 2013). Furthermore,

Evans et al. (2013) confirmed, by thermodynamic analysis, that subunit link regions in the chain resist

to the pulling force generated, an energy that is automatically adjusted over the increasing length of

the growing flagellum (Evans et al. 2013).


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1.2.6 Type IV Secretion Systems

The type IV Secretion System (T4SS) is a long and thin multi-subunit, membrane-spanning

translocation system that is present on the surface of many bacterial species. T4SSs have evolved from

a self-transmissible, single-stranded DNA conjugation system with VirB4-like AAA + ATPase to

systems with an enormous diversity in their overall structure and the types of substrates secreted.

Therefore, this pathway serves to translocate a highly diverse family of substrates, since it has a

functional astounding collective capacity of (Figure 16 A-E):

recognizing and translocating single-stranded (ss) DNA substrates across the membrane to

bacterial or eukaryotic target cell, whereby direct cell contact is required (conjugation

machines);

delivering effector proteins (effector translocator systems) to the cytosol of eukaryotic target

cells in a contact-dependent mechanism;

exporting/importing molecules from/to the extracellular milieu;

delivering killing toxins to bacterial neighbours;

contributing to biofilm development.

Consequently, T4SSs are involved in a variety of functions including pilus formation, toxin

and other protein secretion, gene transfer, and biofilm formation (Grohmann et al. 2018). This type of

secretion system can be found in Gram-positive as well in Gram-negative bacteria and in some

archaea. Furthermore, many pathogenic bacteria use this secretion system as virulence factor and

several intracellular symbionts use it for supporting their colonization and propagation in the

eukaryotic host (Figure 16 E)(Cascales and Christie 2003).

Box5:Flagella Export Apparatus (FEA)

Bacteria are motile microorganisms, to survive they need to swim or swarm depending

on their environment. The bacterial motor that allows this movement it is called the flagellum,

a rotating semi-rigid helical filament. The bacterial flagellum is a highly efficient molecular

nanomachine with approximately 120-140 nm of diameter and about 10 to 20 µm long,

composed by three main structures : the basal body, the hook and the filament, which are made

of about 25 different proteins (Erhardt, Namba, and Hughes 2010). All the components of

these protein complexes are exported, in an unfolded conformation, by the flagellum protein

export apparatus, using ATP and PMF as the energy source.


44

Figure 16. Schematic representation of type IV secretion architecture and functions in bacteria. (A) Conjugative T4SSs translocate DNA from the donor bacterium into various recipients, including other bacteria

or eukaryotic cells. (B) DNA release systems facilitate an exchange of DNA with the extracellular space as well

as biofilm formation. (C) DNA uptake from the environment proceeds by the ComB T4SS. (D) The

Xanthomonas citri T4SS can deliver a protein toxin to kill neighboring G-ve bacterial competitors. (E) Various

pathogenic bacteria and symbionts have evolved T4SSs to deliver effector proteins or DNA–protein complexes

into their host (either eukaryotic target cells or protozoan hosts). The T4SSs can either inject their effectors

directly into the host cell or secrete them into the medium, thereby exerting remarkably different effects on host

cell functions during infection. (F) To achieve genetic exchange during bacterial conjugation, two relaxase

monomers collaborate, adopting distinct structural conformations to provide the two necessary enzymatic

activities for processing the DNA. Adapted from Grohmann et al. 2018)


45

The integrative and conjugative elements (ICEs) are a major form of horizontal gene transfer

mechanism present on bacterial host chromosomal DNA that, when needed, are excised to form a

conjugative DNA circle that can be transferred into other cells (I. Chen and Dubnau 2004; Laverde-

gomez, Sarkar, and Christie 2012). These elements play a key role in bacterial evolution by spreading

genes, which can be involved in cell metabolism, symbiosis, antibiotic resistance, degradation of

pollutants and pathogenesis (Goessweiner-Mohr et al. 2013). ICEBs1 is a ~20.5 kb model ICE for

conjugation in Gram-positive bacteria, found in the B. subtilis genome. It helps to the bacterial

colonization and DNA uptake mechanism (Auchtung et al. 2005; C. A. Lee, Thomas, and Grossman

2012). To the DNA be transfered into the recipient, during the bacterial conjugation, a relaxase

conjugative DNA nucleoprotein complex is delivered by a coupling protein to the DNA translocation

channel. I will now focus and describe in detail the ICEBs1 membrane conjugation machinery required

for this DNA mobilization.

The ICEBs1 T4SS machinery comprises a coupling protein (ConQ), a cell wall hydrolase

(CwlT), an ATPase (ConE) and the membrane channel components (ConB, ConC, ConD, and ConG)

(Leonetti et al. 2015).

The ConQ coupling protein is responsible for the docking of the conjugative DNA to the T4SS

translocation channel and belongs to the FtsK clade of the FtsK-HerA superfamily of ATPases

(Leonetti et al. 2015). ConQ, as the ATPases of its superfamily, forms doughnut-shaped hexamer

with a central channel large enough to hold single or double-stranded DNA (Figure 17), through

its two N-terminal transmembrane helices and a cytoplasmic C-terminal ATPase domain (Iyer et

al. 2004). This protein is essential to the mobilization and conjugation by ICEBs1 (C. A. Lee,

Thomas, and Grossman 2012).

The Gram-positive cell wall is a thick and dense layer. The presence of a cell wall hydrolase is a

key element that allows the correct assembly of the translocation channel, and, allowing the

conjugation. CwlT has two functional domains, a C-terminal domain with DL-endopeptidase

function and an N-terminal domain that works as an N-acetylmuramidase. Mutations in the

catalytic site of CwlT muramidase domain completely abolish the conjugation, while if the

catalytic site of the endopeptidase domain is mutated, it leads to a significant decrease of the

conjugation (Auchtung et al. 2016).

The ICEBs1 membrane channel is a membrane protein complex composed of four integral

membrane proteins ConB, ConC, ConD, and ConG (Figure 17). ConG is the larger component of

this complex, predicted to have seven transmembrane helices, and ConB, as the second larger

component, together may form the bulk of the translocation channel. In contrast, ConC and ConD,

smaller proteins, may work as scaffolding or assembly factors, a role that is still unclear.

ConE is a doughnut-shaped hexamer ATPase, homolog to the Gram-negative canonical VirD4,

which belongs to the VirB4 clade of the FtsK-HerA superfamily, is essential for the DNA

translocation upon the channel (Iyer et al. 2004). The study of its subcellular localization, through


46

fluorescent microscopy using a ConE tagged with the green fluorescent protein (GFP), revealed,

the localization of the ICEBs1 apparatus. In ICEBs1 donor cells ConE-GFP had a polar

localization, and it is possible to see clear dots in the cell pole in close association with the

membrane, while when observed in cells lacking ICEBs1 ConE-GFP was dispersed within the cell

cytoplasm (Berkmen et al. 2010). Leonetti et al. 2015, demonstrated by two-hybrid assays that

ConE directly interacts with ConB, proving that ConB is essential for the recruitment and

maintenance of ConE (Leonetti et al. 2015).

Figure 17. T4SS of ICEBs1. Working model of the ICEBs1 T4SS. This speculative model relies on data from

other T4SSs. The single-stranded conjugative DNA is shown in red, covalently attached to the NicK relaxase.

The presumed coupling protein ConQ likely delivers NicK and associated T-DNA to the membrane-associated

T4SS. ConB and ConG may make up the bulk of the membrane channel. The ConE ATPase may provide energy

for T4SS assembly and/or DNA transfer. Adapted from Auchtung et al. 2016.


47

1.2.7 Holins

The term “holin” or hole formation protein originally refers to phage-encoded pore forming

membrane proteins that control access of phage-encoded endolysins to the peptidoglycan layer. Recent

studies showed that these proteins and their homologues may also be important for a variety of other

functions in Gram-positive phage-free bacteria such as (Saier and Reddy 2015):

spore morphogenesis and germination in B. subtilis;

biofilm formation and DNA release for S. aureus;

programmed cell death and acetate metabolism in S. aureus;

biofilm formation and oxidative stress adaptation in Streptococcus mutans.

Holins are small membrane proteins that play two clear roles in the phage infection cycle.

These proteins can be classified in three different membrane topologies- (I) Class I holins that have

three TMDs, beginning with the N-terminal outside and ending with the C-terminal inside the

cytoplasm; (II) Class II have two TMDs, whereas both ends N- and C-terminal are in the cytoplasm;

(III) Class III just have a single highly hydrophilic TMD, with a disposition of the N- terminal in and

C-terminal out.

The detailed pore formation mechanism, its regulation and the pore dynamics remain

unknown. Studies have shown that until a signal triggers their oligomerization, the small-pore-forming

pin-holins accumulate in the cytoplasmic membrane until it dissipates the PMF. As a consequence this

leads to a membrane permeabilization and large-pore formation that allows the release of endolysins,

which are produced fully folded in the cytoplasm (Figure 18) (Young 2002; Saier and Reddy 2015).

When the endolysins reach the cell wall they cleave various bonds in the peptidoglycan polymer,

depending on the type of endolysin. The genes that encode all the elements involved on this holin-

Box6: Type IV Secretion Systems (T4SSs)

The type IV Secretion Systems (T4SSs) are long and thin multi-subunit, membrane-

spanning translocation systems that are present on the surface of many bacterial species. This

pathway serves to translocate a highly diverse family of substrates, it possesses the collective

capacity to (a) recognize and translocate DNA substrates (conjugation machines) to bacterial

recipients, (b) deliver effector proteins to eukaryotic target cells, (c) exchange DNA with the

milieu, (d) contribute to biofilm development and (e) deliver a killing toxin to bacterial

neighbours. The integrative and conjugative elements (ICEs) are a major form of horizontal gene

transfer mechanism present on bacterial host chromosomal DNA, playing a key role in bacterial

evolution by spreading antibiotic resistance and moving genes, which can be involved in cell

metabolism, symbiosis, degradation of pollutants and pathogenesis. The B. subtilis ICEBs1 T4SS

machinery comprises a coupling protein (ConQ), a cell wall hydrolase (CwlT), an ATPase (ConE)

and the membrane channel components (ConB, ConC, ConD, and ConG) (Leonetti et al. 2015).


48

endolysin system are clustered in a ‘‘lysis cassette’’(I.-N. Wang, Smith, and Young 2000). Therefore,

it is possible to define the holins as “clock” proteins, as they coordinate the timing of the end of the

infection cycle (I.-N. Wang, Smith, and Young 2000).

Real et al. (2005) demonstrated that B. subtilis has a gene, ywcE, which encodes for a holin-

like protein. Performing transcriptional analysis, they prove that this protein is involved in B. subtilis

sporulation (Real et al. 2005). Furthermore, fluorescent microscopy analysis of a strain bearing an

ywcE-gfp translational fusion showed that ywcE localizes at the cell and spore membranes. Its deletion

was also shown to affect the spore morphology. By transmission electron microscopy analysis of the

∆ywcE spores, they indeed showed that the ∆ywcE mutant had a reduced outer coat without the

characteristic striated pattern, and the outer coat failed to attach to the underlying inner coat (Real et

al. 2005).

Figure 18. Schematic depiction of the proteins involved in Gram-negative bacterial cell envelope

disruption by holin-type lysis systems. Pinholins form small heptameric pores that collapse the membrane

potential (the PMF) across the inner membrane, while the more conventional holins form large multisubunit

pores of variable sizes that allow release of fully folded endolysins from the cytoplasm, which in the periplasm

hydrolyze specific bonds in the peptidoglycan cell wall, depending on the lysin type. C+, a cation. Adapted from

Saier and Reddy 2015.


49

1.3 A guarantee of a correct protein translocation in B.

subtilis

As described in the previous chapter, membrane and secreted proteins carry out a multiplicity

of essential functions that directly affect the cellular protein homeostasis, so that the maintenance of

this proteome is of extreme importance for the cell viability. These proteins, from their synthesis until

reaching their final destiny, pass through different structural and folding states. To avoid cell damage

coming from defective and misfolded proteins, which can be caused by different cellular stresses and

leads to an increase of the cytotoxicity level, the cells use specific machineries to overcome these

problems, called quality control mechanism. One part of the quality control strategies is protein

degradation carried out by protease complexes, with the main goal of removing deleterious species and

recycling of amino acids.

Proteases enzymes, that preform protein catabolism by hydrolysis of peptide bonds, are part of

these defense mechanism to eliminate the damaged and misfolded proteins, regulating this way the

fate, activity, localization and protein-protein interactions of many different proteins (Lo and Bond

2009). Some bacterial proteases act as chaperones too, and have a key role in cellular protein quality

control (e.g. B. subtilis WprA feeding protease). Initially, these enzymes were differentiated into two

classes, based on their interaction with the substrate, the endopeptidases, which target internal peptide

bonds, and exopeptidases (aminopepti-dases and carboxypeptidases), the action of which is directed to

the NH2 and COOH terminal domains. Later studies allowed a more detailed classification. Based on

their mechanism of catalysis, proteases can be divided into six distinct classes, aspartic, glutamic, and

metalloproteases, cysteine, serine, and threonine proteases (Lo and Bond 2009). Some bacterial

proteases do not work as individual catalytic devices to hydrolyze the substrate, they have the ability

to interact with each other forming protease complexes. Some of these complexes are crucial, as the

case of the AAA+ proteases (ATPase associated with cellular activities), to the proper functioning and

maintenance of the bacterial cell, as well as to its response to the stress, since these multimeric

machines are able to recognize and degrade misfolded, damaged or aggregated proteins (Dougan et al.

2002).

1.3.1 A bacterial cytoplasmic quality control system: ClpXP protease

Cytoplasmic protease complexes, as ClpXP, are responsible for the degradation of defective or

denatured membrane or secretory proteins that, due to stress conditions, cannot be translocated into or

out of the membrane and are arrested in the cytoplasm. ClpXP consists of two subunits, the ClpX

subunit, which has ATPase activity and form a hexameric ring, and the ClpP, which is a double-

heptameric serine protease (Figure 20). The interaction between these proteins was observed for the

first time in the early 1990s, Gottesma et al. named and identified E. coli ClpX as the protein that, in

the presence of ClpP, induces the ATP-dependent degradation of λ0 protein. They showed that the


50

clpX and clpP genes form an operon and are co-transcribed in a single heat-induced mRNA

(Gottesman et al. 1993). As for E. coli ClpP, it has been demonstrated that B. subtilis clpP is also

induced by heat shock stress. Additionally, it was shown that B. subtilis clpP is induced by salt and

ethanol stress, as well as after a puromycin treatment. Indeed after such a stress exposure the amount

of clpP specific mRNA increased; however the highest increase was shown after heat shock stress

(Gerth et al. 1998). Therefore, these experiments suggested that B. subtilis ClpP is involved in the cell

response to puromycin treatment or heat shock stress, through the degradation of abnormal proteins

resulting from the stress. It was also demonstrated that the deletion of clpP or clpX in B. subitilis

grown in stress conditions, induced the production of elongated cells (Figure 19) and an impaired

growth, indicating that B. subtilis ClpXP have substrates involved in cell morphology or cell division

and emphasized the importance of ClpXP as proteases or chaperones in stress tolerance (Gerth et al.

1998).

Figure 19. Scanning electron micrographs of B. subtilis IS58 (A), the clpP (B) and clpX (C) mutant strains

grown exponentially. Adapted from Gerth et al. 1998.

ClpP exhibits a cylindrical structure composed of two associated heptameric rings and, as a

typical serine protease, contains internal (inner surface) proteolytic sites composed of the conserved

catalytic triad (Ser-His-Asp) (B. G. Lee, Kim, and Song 2011). The hexameric ClpX ATPase ring

binds to the heptameric ClpP ring in a symmetry mismatch, aligning the two axial pores that creates a

continuous central channel between them (Figure 20). Allowing the translocation, of the denatured

and linear protein substrate, from the narrow ATPase pore into the concealed proteolytic chamber

(Olivares, Baker, and Sauer 2015). Despite the fact that high-resolution structures of the ClpXP

complex do not exist, it was demonstrated, by different studies and techniques, that the stability of this

complex relies on interactions involving flexible loops near the axial pores of each ring and another set

between peripheral structural elements, as represented on Figure 21 (Baker and Sauer 2012). Joshi

and colleagues demonstrated that the major determinant of interaction within E. coli ClpXP complex,

are surface ClpX loops, which contains a IGF conserved motif. They observed mutations that ClpX

IGF loop lead to a weakening or elimination of the ClpX-ClpP interaction without affecting other


51

ClpX functions (Joshi et al. 2004). The IGF motif is present in B. subtilis ClpX, however these

structural interactions have not been proved yet.

Figure 20. Cartoon model of substrate recognition and degradation by the ClpXP protease. In an initial

recognition step, a peptide tag in a protein substrate binds in the axial pore of the ClpX hexamer. In subsequent

ATP-dependent steps, ClpX unfolds the substrate and translocates the unfolded polypeptide into the degradation

chamber of ClpP for proteolysis, where it is cleaved into small peptide fragments. (Baker and Sauer 2012).

Figure 21. Interaction of ClpX and ClpP. The ClpXP complex is stabilized by peripheral interactions between

the IGF loops of ClpX and hydrophobic clefts on ClpP, and by axial interactions between the pore loops of ClpX

and the N-terminal stem-loop of ClpP (Baker and Sauer 2012).


52

The interaction between this two proteins is essential, as the ClpX protein is responsible for

recognition and binding of the substrates, and it allows the degradation of larger protein substrates as

for example disulphide-bonded substrates which require the translocation of three polypeptide chains.

The crystal structure of the ClpX structure revealed a small pore almost completely closed, that will

need to expand and adapt to the different substrate sizes (Baker and Sauer 2012; Gersch et al. 2015).

So, how does the ClpX narrow pore expand according to the substrate size?

Each ClpX protomer has two domains, the large and the small domains, that assemble,

through the binding of the small domain of one protomer to the large domain of the next one, in an

asymmetric hexamer that can bind a maximum of four molecules of ATP and/or ADP (Glynn et al.

2009). Due to its asymmetry and domain-domain orientation the ClpX hexamer can be divided into

subunits of two main classes, the “loadable” or L subunits and the “unloadable” or U subunits (Glynn

et al. 2009; Baker and Sauer 2012). In the L subunit, the large and small AAA+ domains of each

protomer connect in a nucleotide-binding conformation, corresponding to four subunits of the

hexamer. Interestingly, on the two U subunits can occur a rotation of 80° between the bound domains

of the protomers (large and small AAA+ of each protomer), originating movements as large as 30 Å

that destroy the nucleotide-binding site (Glynn et al. 2009; Baker and Sauer 2012). Consequently, the

two U subunits, create two flexible sides of the hexamer that allow the adaptation of the pore to

different substrate sizes, extending the pore to accommodate larger substrates and then refold and

shortening it to contact with smaller substrates (Glynn et al. 2009; Baker and Sauer 2012). This

movement can be compared to the unhinging of the jaws of a snake, which allows it to swallow large

prey, as it is demonstrated on Figure 22.

Figure 22. Representation of the expansion and contraction of the axial pore by a snake-jaws model. The

main contacts between these jaws are formed by the interfaces between the large and small AAA+ domains of

the unloadable (U) ClpX subunits. In which pore size is controlled by the size of the substrates and the

conformation of the hinge region and flanking structure of the unloadable (U) ClpX subunits. Opening of these

interfaces, as shown in the exaggerated right cartoon, provides a potential mechanism for pore expansion to

accommodate large substrates, including those with multiple chains. Each colour represents a ClpX protomer.

(L) loadable subunits. (Baker and Sauer 2012).

This mechanism starts by the recognition of substrates by ClpX, through the binding to short

unstructured peptide sequences called degradation tags or degrons, which are typically located at the


53

N-terminal or C-terminal end of a target protein, or to adaptor proteins, that modulate and enhance the

substrate recognition by the respective ATPase partner (Dougan et al. 2002; Kirstein et al. 2006). The

adaptor proteins bind to specific substrates and form a delivery complex with the ATPase partner, by

binding to its N-terminal domain (Kirstein et al. 2006; Molière and Turgay 2009; Elsholz, Birk, and

Charpentier 2017). The ClpX adaptor proteins of B. subtilis are YjbH and CmpA (Kirstein et al. 2009;

Elsholz, Birk, and Charpentier 2017).

YjbH is a cytosolic protein involved in sensing various stresses and is an adaptor protein for

the ClpXP catalyzed degradation of Spx factor, being, this way, involved in the resistance against

nitrosating agent. The name of transcription factor Spx was originated from the identification of

genetic mutations on its encoding gene (yjbD), that caused a suppression clpP or clpX (Nakano et al.

2001). High concentrations of Spx were observed in the clpP mutant, higher than in the wild-type

strain, indicating that ClpP is necessary to maintain the Spx concentration low (Nakano et al. 2001).

YjbH exposes the Spx degradation tag, the ClpX recognition site, mediating its degradation. This

recognition site, part of the Spx 28-amino-acid C-terminal segment, is buried by its own structure

buries, a structure that is disrupted by YjbH interaction, thereby promoting Spx degradation by ClpXP

(Garg et al. 2009; Chan, Hahn, and Zuber 2014). This way, under normal conditions, the YjbH-ClpXP

interaction suppress Spx activity through its degradation, while under stress conditions, e.g. heat or

oxidative stress, leads to the YjbH aggregation that promotes the stabilization and accumulation of

Spx (Chan, Hahn, and Zuber 2014; Elsholz, Birk, and Charpentier 2017).

On the other hand, CmpA adaptor protein, together with ClpXP, are involved in the quality-

control pathway that maintain the integrity of the spore envelope, by removing defective sporulating

cells (Tan et al. 2015; Elsholz, Birk, and Charpentier 2017). Under correct sporulation conditions, the

cells produce spores with a proper envelope, ClpXP degrades CmpA and the sporulation continues.

However, in those cells harbouring defects or misassembled spore envelopes, CmpA stabilizes and

delivers the coat morphogenetic protein SpoIVA for degradation by ClpXP, that, consequently,

destabilizes the forespore and leads to cell lysis, thereby eliminating the defective cells and ensuring

the correct spores assembly within the population (Tan et al. 2015; Elsholz, Birk, and Charpentier

2017).

This structural and activity coordination between the six-fold symmetric ClpX ATPase and

seven-fold symmetric ClpP peptidase allows the unfolding of bound substrates in an ATP consuming

reaction and the transportation of the unfolded polypeptide into the ClpP proteolytic chamber, that are

consequently degraded to small peptides that exit the chamber by diffusion, as represented at Figure

20 (Glynn et al. 2009; Baker and Sauer 2012; Elsholz, Birk, and Charpentier 2017).


54

1.3.2 Membrane-bound proteases, an important quality control step for

membrane and secretory proteins.

The bacterial cell envelope, membrane and cell wall, the external layer of bacterial cells, is very

important for the integrity and viability of bacteria, as it provides physical protection, determines the

cell shape and is the principal stress-bearing element (Scheffers and Pinho 2005; Thomas, Daniel, and

Suzanne 2010). These are sophisticated, dynamic and complex structures that, which in addition to

protecting, allows selective passage of nutrients from the outside, translocate secretory proteins and

waste products from the inside. Gram-negative bacteria cell envelope, such for E. coli, is composed of

two membranes, the cytoplasmic membrane and an outer membrane, and between these structures a

membrane-enclosed periplasm, a dynamic and metabolically highly active environment. The outer

membrane of Gram-negative bacteria contains lipopolysaccharides in its outer leaflet and

phospholipids in the inner leaflet.

In contrast, Gram-positive bacteria have a cytoplasmic membrane and a cell wall. The cell

wall is a complex structure composed of surface proteins, teichoic acids and a thick layer of

peptidoglycan (PG). Peptidoglycan, also called murein, is a heteropolymer composed of long glycan

chains, made up of alternating β-1,4-linked N-acetylglucosamine (GlcNAc) and N-acetylmuramic acid

(MurNAc) subunits, which are cross-linked by peptide bridges to form a strong but flexible structure

(ref). Peptidoglycan is present in almost all bacteria, in B. subtilis these glycan strands are considered

to be long (between 50 and 250 disaccharide units) when compared to, for instance, Staphylococcus

aureus strands (between 3 and 10 disaccharide units), except in Mycoplasma and a few other species

that lack detectable cell walls (Vollmer, Blanot, and De Pedro 2008). To recoup the absence of

periplasm, Gram-positive bacteria are able to lipo-modify homologues of periplasmic proteins and

attach them to the outer leaflet of the cytoplasmic membrane, as well as retain some secretory proteins

in the cell wall by ionic or covalent attachment to the peptidoglycan (Sarvas et al. 2004; Schneewind

and Missiakas 2014). This way, the Gram-positive cell wall can be described as a thicker but dynamic

and flexible structure rich in cell surface proteins, including quality control proteases,

extracytoplasmic chaperones, autolysins, surface layer proteins, and substrate binding proteins (Pohl

and Harwood 2010a; Schneewind and Missiakas 2014). In addition to peptidoglycan, an important

class of cell surface glycopolymers in Gram‐positive bacteria are the phosphate rich teichoic acids

(TAs). These molecules play a role in a large variety of functions, such as in maintaining the

physicochemical properties of the cell surface, cation homeostasis, resistance to antimicrobial peptides

and lytic enzymes, acting as phage receptors, in cell division, biofilm formation and host adhesion.

There are two types of TAs, distinguished by the way they are covalently linked to the surface, the

lipo-teichoic acids (LTAs), which are anchored to the cytoplasmic membrane, extending from the cell

into the peptidoglycan layer, and the wall teichoic acids (WTAs), which are covalently attached to the

peptidoglycan layers and extend beyond them. Together, the LTAs and the WTAs, create a negative


55

gradient that goes from the bacterial cell surface until the outer most layers of the PG (C Weidenmaier

and Peschel 2008; Pasquina, Santa Maria, and Walker 2013). B. subtilis has two distinct phosphate

rich WTAs, the poly(glycerol phosphate) and poly(glucosyl N-acetylgalactosamine 1-phosphate),

although the last one is present in minor amounts (Freymond et al. 2006). These anionic polymers and

the lipo-teichoic acids create a high density of negative charge in the cell wall, an environment into

which secretory proteins emerge in an unfolded state from the translocase (Hyyrylainen et al. 2000).

As described before in this introduction the Gram-positive architecture is compatible to

protein export, a closer look at the cell envelope reveals that it offers a very challenging environment

for proteins. From the correct insertion of membrane proteins and their maintenance to the unfolded

proteins that emerge from the translocase, which should fold into their native configuration quickly

without forming intra or intermolecular interactions that could block the translocation machinery

and/or block cell wall synthesis sites, either or which would compromise cell viability. This reinforce

the importance of having membrane quality control systems. As is the case of HtrA, HtrB and WprA

for the Sec pathway, as they are serine proteases, belonging to the quality control protease family,

combating the extracellular folding stress, and preventing potentially fatal obstruction of the secretory

translocase. Another serine protease of the Sec pathway is the Signal peptide peptidase A (SppA), that

will be described in the next subchapter.

1.3.3 Signal peptide peptidase A (SppA)

As detailed in the second chapter of this introduction the rapid removal and recycling of the

cleaved signal peptide is crucial for maintaining a proper protein translocation, an action engaged by a

membrane protease named signal peptide peptidase. The E. coli signal peptide peptidase, encoded by

the gene sppA, was purified and characterized in 1987 by Ichihara and his colleagues (Ichihara et al.

1986; Suzuki et al. 1987). They showed that this protein forms a tetramer, in which each protomer is

estimated to have 67 kDa, and it is able to digest, in the presence of the cell envelop, the accumulated

signal peptides and non-specifically other proteins (Ichihara et al. 1986).

Through sequence comparison between E. coli and B. subtilis genomes, a homologue of the E.

coli sppA gene was found in B. subtilis, the yteI. YteI was renamed signal peptide peptidase A (SppA)

of B. subtilis as E. coli SppA (SppAEC), since these two proteins have 49% of sequence similarity

(Bolhuis et al. 1999). In contrast, B. subtilis SppA (SppABS) is a smaller membrane protease with

about 36 kDa that assembles as an octamer, which protomer have a globular and extension part (Nam,

Kim, and Paetzel 2012).

As characteristic of serine proteases, the SppA catalytic domain consists in a dyad of a serine

and a lysine (Nam, Kim, and Paetzel 2012). In order to improve the structure resolution, SppAEC and

SppABS, were purified without the N-terminal transmembrane region and crystalized. The SppAEC

protomer structure showed that the active site residues are in opposite domains, the lysine general base

(K209) was identified in the N-terminal domain while the serine nucleophile (S409) is in the C-


56

terminal domain, which means that they just interact with the chains are side-positioned (Kim, Oliver,

and Paetzel 2008; P. Wang et al. 2008). Differently, the crystal structure of SppABS revealed that a

serine (S147) residue, located at the globular region of the protomer, and a lysine (K199), located at

the extension region, are quite distant from each other, as shown on Fig.23. Demonstrating that

SppABS on its own is not able to catalyse a substrate, as the active-site catalytic dyad is promoted by

the interaction between the S147 of one promoter and the K199 of the adjacent protomer (Nam, Kim,

and Paetzel 2012).

Figure 23. The B. subtilis SppA protein. (A) A schematic diagram showing the full-length SppA B. subtilis

with its predicted transmembrane segment (pink) and the confirmed amino- terminal thermolysin cleavage site.

The light-green region is what is observed in the electron density. (B) A topology diagram of SppA B. subtilis

showing the full-length protein with β-strand as arrows and α- helices as cylinders. The protein is colored as a

gradient from N- terminus (blue) to C-terminus (red). The regions not seen in the electron density are shown as

broken lines. (C) A cartoon diagram showing the tertiary structure of the SppA B. subtilis protomer. The β-

strands are shown as arrows, and the α- helices are shown as cylinders. The color scheme is the same as in (B).

The nucleophile Ser147 (red) and the general base Lys199Ala (blue) are shown as spheres (Nam, Kim, and

Paetzel 2012).

A

B

C


57

Figure 24. Comparison between octameric SppABS and tetrameric SppAEC. (A) Schematic representation of

SppAEC tetramer. (B) Schematic representation of SppABS octomer. The catalytic dyads are shown (general base,

blue spheres; nucleophile, red spheres). One protomer in each oligomer is shown in black in (A) and (B). (C) A

cartoon diagrams of an SppA E. coli protomer: the C-terminal domain is colored darker gray, and the region

linking the domains is colored yellow. (D) A cartoon diagrams of an SppA B. subtilis protomer.

Both oligomeric forms of SppAEC and SppABS assemble into a dome-shaped structure (

Figure 24), with a large opening of the bowl, created by the globular region, facing the outer

leaflet of the cytoplamic membrane and a axial hole (small hole), originated by the extension region,

facing the periplasm or extracellular media, respectively (Kim, Oliver, and Paetzel 2008). The interior

of the dome is predominantly hydrophobic and it has an inside midway up a continuous concave

groove that encircles the entire inner bowl, containing the active sites and the substrate binding

pockets (Kim, Oliver, and Paetzel 2008; Nam, Kim, and Paetzel 2012). The substrate binding pockets

of SppABS were described as deeper and wider than the SppAEC pockets, as it is represented on the

cross plan of Figure 25. Although the substrates, E. coli and B. subtilis signal peptides, are predicted

and experimentally shown to have a similar residues content the signal peptides of Gram-negative

bacteria are in general shorter than the Gram-positive ones (Nam, Kim, and Paetzel 2012).

In 2013, Nam and colleagues, revealed an important step of SppABS activity regulation

through its crystal structure. They solved the crystal structure of the SppABS∆2-54

deleted of the first 54

amino acids, corresponding to the N-terminal transmembrane domain, and of an active-site mutant


58

(K199A) of SppABS∆2-54

, which is an inactive form of SppA. The structure of the SppABS∆2-54

inactive

form revealed the presence of the C-terminal region bound to the catalytic groove, meaning that

SppABS∆2-54 self-processes its own C-terminal (Nam and Paetzel 2013). They also demonstrate that this

C-termini peptide possibly have a role in the regulation of proteolysis, as it is able to compete with a

synthetic peptide, based on the bound peptide, for the SppABS active sites (Nam and Paetzel 2013). In

agreement with the information obtained for SppAEC, in vitro studies demonstrated that SppABS not

only digests the signal peptides as it is able to digest fully folded proteins. An active form of purified

SppABS was able to digest overnight a folded E. coli lipoprotein (BamD) involved in outer membrane

protein assembly (Nam, Kim, and Paetzel 2012).

Figure 25. Cross section representation of the oligomeric dome shape structure of SppAEC and SppABS. (A)

A section through the middle of SppAEC revealing the dimensions of the opening to the bowl-shaped structure.

The catalytic residues S409 (green) and K209 (magenta) are shown as van der Waals spheres. (B) A cross-

section view of the SppABS octamer, revealing the continuous concave groove that encircles the entire inner

bowl, with dimensions given. Adapted from Kim, Oliver, and Paetzel 2008 and S. E. Nam, Kim, and Paetzel

2012.

Furthermore the SppABS function and role in secretion was studied by Bolhuis et al. in 1999,

where was shown that the deletion of SppA induced an increase of pre-AmyQ accumulated in the

cytoplasm and, consequently, a slight decrease in the secreted mature AmyQ, a normally highly

secreted protein. Suggesting, this way, that SppA has a role on the processing of the pre-proteins under

hyper-secreting conditions. B. subtilis have extracytoplasmic function σ factors (ECF) σM, σW and σX

that induce a stress response against lantibiotics, contributing for the resistance to this antibiotic

peptides. SppABS is expressed under the control of a σA factor (a constitutive expression through the

different growth phases) and also under the control of σM, which can indicate possible connection to

stress response and lantibiotic resistance (Huang and Helmann 1998; Nicolas et al. 2012). The

possible involvement of SppABS in cleaving and consequently protecting the cell against peptide

antibiotics that insert themselves into the cytoplasmic membrane was investigated by Kingston and

colleagues (Kingston, Liao, and Helmann 2013a). Through diffusion disks assays, they demonstrated

that the deletion of SppA decreases the strain sensitivity to nisin, lantibiotic that binds to lipid II and


59

inhibiting cell wall synthesis and forming pores in the cytoplasmic membrane, which suggests a

possible role on antibiotic cell protection (Kingston, Liao, and Helmann 2013a).


60

1.4 Scope of the Thesis

Around 5-10% of the bacterial proteins produced are transported from the cytoplasm to the

extracellular compartment to perform several important functions, such as the provision of nutrients,

cell-to-cell communication, and release of toxins and detoxification of the environment. In B. subtilis,

the major route for the export of these proteins involves mainly the highly conserved Sec pathway

(Harwood, C. R. and R. Cranenburgh 2008; Tjalsma, H., et al. 2004).

The Sec pathway, described in the introduction section 1.2.1 of the introduction, mediates the

translocation across the hydrophobic membrane barrier of secretory and membrane proteins in an

unfolded conformation. Briefly, this secretion process can be divided into three steps: targeting,

translocation and release. First the secreted proteins are synthesised as precursors (pre-pro-protein)

with a N-terminal signal peptide (SP), which is recognized by cytoplasmic factors (Signal Recognition

Particle- SRP) or chaperones (CsaA) involved in targeting the pre-protein to the translocon SecYEG.

The translocation of these proteins across the channel is driven by the translocon motor SecA, an

ATPase that interacts with the substrate proteins and binds to the SecYEG complex. The repeated

cycles of binding, hydrolysis of ATP, conformational changes of a dimeric SecA leads to the

translocation of the secretory protein through the SecYEG complex. Subsequently, the signal peptide

is cleaved resulting in the release of the protein from the membrane. Finally, the protein undergoes a

maturation process carried out by other accessory proteins, such as the foldase PrsA, or the Bdb thiol-

disulfide oxido-reductases (Driessen and Nouwen 2008; Kang et al. 2014; H Tjalsma et al. 2004).

Although these mechanisms involved in the Sec pathway has been extensively studied in B.

subtilis, the knowledge about the protein complexes is limited. For example, it was shown, in E. coli,

that the holotranslocon is a large complex where the YidC and SecYEG are linked by a subcomplex

consisting of YajC and SecDF. SecDF regulates the interaction and activity of SecA with SecYEG.

Physical interactions, organization, activity and bioenergetics of these two sub-complexes SecYEG-

YidC and SecDF-YajC were studied in vivo and in vitro (Duong et al. 1997; Schulze et al. 2014).

Although paralogous proteins have been identified in B. subtilis, such detailed data on these

complexes are not available. This raises questions about the organisation of the holotranslocon in B.

subtilis.. In the first chapter of results, we will show the efforts made to detect physical interactions

involving the B. subtilis (holo)translocon proteins as well as the possible interactions between the

accessory proteins involved in the maturation of secreted proteins. The goal was also to

discoverunknown protein partners interacting with these complexes. In this chapter we showed a

possible protein complex between a known protease, called SppA, and an unknown protein YteJ.

The chapter 2 of the results is dedicated to the study of the membrane complex SppA/YteJ.

The signal peptide peptidase A (SppA), a component of the Sec machinery of B. subtilis, was named

as such because sequence comparison with that of E. colirevealed that these two proteins share 49% of

sequence similarity (Bolhuis et al. 1999). SppA is a membrane bound serine protease that forms a


61

homo-octameric dome-shaped complex described to be involved in the recycling of the remaining

signal peptides (SPs), cleaved off the translocated proteins, in the membrane. SppA is also involved in

the resistance to nisin, an antibacterial peptide of the lantibiotic family (Bolhuis et al. 1999; M. Cao et

al. 2002; Nam and Paetzel 2013). Its function and role in protein secretion were studied by Bolhuis et

al. (1999), who showed that the deletion of SppA induced a mild decrease in the secretion of mature

secretory proteins, suggesting that SppA has a role on the processing of the pre-proteins under hyper-

secreting conditions (Bolhuis et al. 1999). However, Saito et al. (2011), showed that the disruption of

E. coli sppA gene did not affect in vivo cleavage of the SPs and that the B. subtilis RasP, a S2P

protease, is involved in proteolysis of SPs of some secretory proteins and. Recently in 2017, Neef et

al. gave a proof that the overexpression of RasP in B. subtilis improved the secretion of an

heterologous and highly secreted AmyAc amylase by tenfold, proving that the overexpression of a

bacterium membrane protease can boost the protein production (Neef et al. 2017). These results

changed the view on the SppA function, rising new questions about this topic. In B. subtilis sppA gene

is in an operon with another gene called yteJ. From its sequence, the protein YteJ, of unknown

function, is predicted to be an integral membrane protein belonging to the RDD family, a family of

proteins of unknown functions. To better understand the influence of YteJ on SppA function and to try

to clarify SppA role, we divided this study in two main parts – the in vivo and in vitro study. In vivo,

we tried to uncloak the main role of the SppA/YteJ complex in the cell, starting to study the effect of

the deletion of these two genes, their regulation by two sigma factors (σA and σW), their cellular

localization by fluorescent microscopy, the involvement of both SppA and YteJ in the lantibiotic

resistance.

63

RESULTS

65

2 RESULTS

2.1 Identification of membrane protein complexes involved

in the B. subtilis Sec pathway

2.1.1 SPA-tag constructions

In order to study the interactions within the B. subtilis (holo)translocon, as well as the possible

interactions between the accessory proteins or unknown protein partners interacting with these protein

complexes, we constructed strains harbouring the 3'-translational SPA sequence fusions in the genetic

background of the BSB1 strain. This strain is tryptophan-prototrophic (trp+) derivative of the 168

trpC2 strain as described in (Nicolas et al. 2012).

A set of proteins of the Sec secretion pathway (see Table 1 below) was chosen to be SPA-

tagged for tandem affinity purification. For that purpose, the 3' ends of the corresponding genes were

fused to the SPA-tag sequence present in the plasmid pMutin-SPA-LIC. The constructed plasmids

were used to transform the BSB1 strain by a Campbell-like integration. Moreover, the pMutin-SPA-

LIC is an integrative plasmid that contains an IPTG inducible / LacI repressible Pspac promoter that,

once integrated in the chromosome, is just upstream of the downsteam genes to ensure their

expression.

The construction of these plasmids had some setbacks: it had been only possible to obtain nine

strains out of ten, as PrsA-SPA strain was not obtained. PrsA is an essential protein for extracellular

folding of some secreted proteins after their translocation. The addition of the SPA-tag might have

impaired its activity, therefore leading to an unviable strain.

Table 1. Set of proteins involved in the Sec pathway chosen to be SPA-tagged for tandem affinity

purification.

Name Protein Function Molecular weight

(kDa)

SecDF

Preprotein

translocase subunit

(ATPase)

Regulates the

interaction and

activity of SecA with

SecYEG

81.47

SecG Sec tanslocase

subunit Protein secretion 7.86

SipT Signal peptidase I

Cleavage of the

signal peptide from

the emerging protein

21.71

Chapter 2.RESULTS

66

SipS Signal peptidase I

Cleavage of the

signal peptide from

the emerging protein

20.90

SppA Signal Peptide

Peptidase

Recycling of the

cleaved signal

peptide

36.52

YteJ Stress protein Unknown function 18.98

SpoIIIJ Membrane protein

translocase

Insertion of

membrane proteins

and protein secretion

29.37

BdbB Thiol-disulfide

oxidoreductase

Oxidative folding of

proteins 17.00

BdbC Thiol-disulfide

oxidoreductase

Oxidative folding of

proteins 15.57

PrsA Post-translocation

molecular chaperone Protein Folding 32.36

The final strains harbouring the 3'-translational SPA fusions are listed in Table S2. Growth

curves (Figure 26) and Western-blot (Figure 27) analyses were performed to check the possible

growth defect due to the constructs and the production of the corresponding tagged proteins

respectively. Strains were grown in Luria Bertani (LB) medium and the OD was recorded. The growth

of none of the strains was affected by the SPA-tag fused sequence to any of the proteins chosen

(Figure 26). To check the expression of the SPA-tagged proteins in these strains by Western-blot,

samples of cells were taken in exponential, transition and stationary phases. Although the nine strains

harboured the 3'-translational SPA sequence fusions to the selected genes, only six proteins (SecDF,

SecG, SpoIIIJ, YetJ, BdbB and BdbC) were detected by Western-blot using anti-FLAG anti-bodies

(Figure 27).

The SipS-SPA, SipT-SPA and SppA-SPA were not detected (Figure 27 (B)-C and (C)-D, F).

However, for these three proteins, the C-terminal domain (fused to the SPA-tag in our constructs) is

predicted to be located on the extracellular side of the cytoplasmic membrane, while for all the other

protein fusions detected successfully the C-terminal domain is predicted to be inside the cytoplasm.

Therefore, it might be possible that the SPA-tag sequence was cleaved by extracellular proteases when

the three proteins are translocated and inserted into the membrane. To study these three proteins we

decided to construct strains harbouring a N-terminal SPA-tag fusions instead of the C-terminal fusion.

A SPA-tag sequence was fused to the 5' end of the corresponding full-length genes and cloned into

another plasmid, pSG-SPANter, which allows the integration of the fused gene at the amyE locus by

67

homologous recombination. Unfortunately, although several attempts were done, these constructions

could not be obtained.

Figure 26. Growth analysis of strains harbouring C-terminal SPA-tag fusions. The different strain cultures

were grown overnight in LB supplemented with 0.6 µg/ml Erythromycin and 0.5mM IPTG at 37°C, and in the

next day diluted 1:200 into a new medium with the same supplements. Compared to the BSB1 WT strain the

SPA-tag fusions do not affect the growth of the mutant strains. Samples were taken on specific time points.

SecDF-SPA was detected (Figure 27(B)-A) as a main band of the right molecular weight in

the three growth phases, with additional bands corresponding to proteins of lower molecular weight,

probably products of SecDF-SPA degradation.

SecG-SPA was detected (Figure 27(B)-B) just in transition and stationary phase with the

lower apparent expression level compared to that of the other SPA-tagged proteins of our analysis.

BdbC-SPA (Figure 27(C)-E), YteJ-SPA (Figure 27(D)-G), SpoIIIJ-SPA (Figure 27(D)-H)

and BdbC-SPA (Figure 27(D)-H) were also detected in the three growth phases, showing three

intense bands. These results showed the six detectable SPA-tagged proteins (SecDF, SecG, SpoIIIJ,

YetJ, BdbB and BdbC) could then be purified using this tandem affinity purification technique.

Chapter 2.RESULTS

68

Figure 27. Expression of the SPA tag fused proteins. (A) Table with the corresponding molecular weights of

each SPA tagged protein in kDa. (B), (C) and (D) Western blot analysis, using an anti-FLAG specific

antibodies. Cell samples from exponentional, transition and stationary phases were loaded for each strain:

SecDF-SPA (a), SecG-SPA (b), SipS-SPA (c), SipT-SPA (d), BdbC-SPA (e), SppA-SPA (f), YetJ-SPA (g),

SpoIIIJ-SPA (h) and BdbB-SPA (i).

69

2.1.2 Purifications of the SPA-tagged proteins

The tandem affinity purification approach is based on two sequential affinity purifications as

the SPA-tag is composed of two affinity tags (FLAG and CaM peptide). This double affinity

purification allows the elimination of more contaminants compared to a single affinity purification.

This technique is also use in native conditions to purify protein complexes, i.e. without using cross-

linkers that sometimes induce “false” interactions. The C-terminal SPA-tagged proteins (SecDF,

SecG, SpoIIIJ, YteJ, BdbB and BdbC) that were proved to be well expressed (see western-blot Figure

27) were purified using this procedure. For the identification of the co-purified proteins by liquid

chromatography coupled to tandem mass spectrometry (LC-MS/MS), the purified samples were

loaded on SDS-PAGE for a short migration. Large bands (0.5 cm), which comprises all the co-purified

proteins, were cut out of the gel and were sent for analysis by LC:MS/MS.

According to Schulze et al. (2014), SecDF, an ATPase, and YidC, an membrane protein

translocase, form a subcomplex that interacts with the SecYEG in E. coli, and all together constitute

the holo-translocon (Schulze et al. 2014). Therefore, we expected that SecDF and SpoIIIJ, respective

B. subtilis homologous of E. coli SecDF and YidC, form a sub-complex in B. subtilis as well. As such,

we expected to identify and detect, by LC-MS/MS, these proteins together in the same sample after the

SPA-purification of SecDF-SPA or SpoIIIJ-SPA. However, SecDF was not found in the co-purified

proteins of the SpoIIIJ-SPA purified sample, and conversely, SpoIIIJ was not detected either in the

SecDF-SPA purified sample (data not shown). These results showed that, as opposed to E.coli

(Schulze et al. 2014), SecDF and SpoIIIJ do not co-purify in B. subtilis. Perhaps, in B. subtilis this two

proteins form a weak interaction that could be disrupted by the presence of a SPA-tag. Although

counter-intuitive, the difference could come from the detergent (n-Dodecyl β-D-maltoside- DDM)

concentration used to solubilize the membranes, since in E. coli 2% DDM was used (Schulze et al.

2014) and our experiments were performed with 1% DDM. In B. subtilis, these complexes might be

more susceptible to the DDM concentration and even 1% is too high so that the weak interactions

might be abolished. In order to test this hypothesis, we used four different concentrations 0.5%,

0.75%, 1% and 2% of DDM to solubilize the B. subtilis membranes from the SecDF-SPA and SpoIIIJ-

SPA strains. To maximise the chances to get these two proteins co-purified in at least one condition,

we also used only one affinity chromatography, i.e. the calmoduline resin. After purification in these

different conditions of DDM concentrations, to check the quantity levels of the captured proteins,

SpoIIJ-SPA or SecDF-SPA, we analysed each purified sample by Western-blot (Figure 28 (A)).

These results showed that the purification of both proteins were more or less the same whatever the

different concentrations of DDM (Figure 28 (A)). The LC-MS/MS results corresponding to the

analysis of these samples, that have identified numerous proteins (mainly contaminants), are focused

only on the presence or absence of the other possible partners (SpoIIIJ for SecDF-SPA and SecDF for

SpoIIIJ-SPA) and are showed as two tables in Figure 28 (B). These results showed that the SpoIIIJ

Chapter 2.RESULTS

70

was not detected in the SecDF-SPA purified samples. However, low amounts of SecDF were detected

in all the SpoIIIJ-SPA samples. Altogether, these results showed that SecDF and SpoIIIJ do not form a

stable complex in B. subtilis in the conditions tested. The small amounts of SecDF found in the

SpoIIIJ-SPA purified samples might be an indication of a weak interaction between the two proteins

or a simple contamination as only one affinity column was used. In either cases we could conclude

that these two proteins form a stable complex, at least in our tested conditions.

Figure 28. Purification of SpoIIIJ-SPA and SecDF-SPA protein with different concentrations of DDM. (A)

Western blot analysis, using an anti-FLAG specific antibodies, of SecDF-SPA and SpoIIIJ-SPA in the presence

of a range of n-Dodecyl β-D-maltoside (DDM) concentrations, from 0.5% up to 2%. (B) Identification by LC-

MS/MS of the expected protein partner: poitive (٧) negative (-).

71

The other experiments using SPA-tagged proteins (SecG, BdbB, and BdbC) did not provide

evidence for the existence of any stable complexes involving proteins known to be part of the

translocon complex.

The only exception came with the finding of SppA, the so-called signal peptide peptidase, that

co-purified with YteJ-SPA. This result was expected as preliminary results obtained in our lab from

Blue native PAGE (BN-PAGE) experiments had indicated that these two proteins might form a

complex. Two tandem affinity purifications of the YteJ-SPA protein and LC-MS/MS analysis showed

that the relative quantities of the two proteins could be inferred from the LC-MS/MS data. The

SppA/YteJ ratios found were 2:1 and 1:1. As both genes are in operon, it can be expected that the two

proteins are translated in a 1:1 ratio, which would be in accordance with our experimental results.

2.1.3 Optimisation of the BN-PAGE technique

The BN-PAGE (for Blue Native PAGE) can be used to isolate protein complexes

from the membrane fraction, to determine the molecular weight of native protein complex, to obtain

information about their oligomeric states and to identify physiological protein–protein interactions

(Wittig, Braun, and Schägger 2006). We therefore used the BN-PAGE as a complementary technique

to identify the components of membrane protein complexes involved in the Sec pathway. As this

electrophoretic technique is performed in native conditions, theoritically, the different protein

components of a stable complex are not separated and migrate together in the gel. The n-Dodecyl β-D-

maltoside (DDM) was the detergent chosen to solubilize the membrane samples obtained after

ultracentrifugation of disrupted B. subtilis cells. After solubilisation (1% DDM) the samples are

supplemented with an anionic dye, Coomassie blue G-250, and the protein complexes are separated

according to their sizes by electrophoresis on a gradient 4-16% polyacrylamide gel. High molecular

weight complexes are running high in the gel. If the concentration of detergent is too high, these

complexes dissociate and individual proteins are running lower in the gel.

We used the SecDF-SPA strain to analyse if the BN-PAGE can used for our study as SecDF-

SPA is well expressed and can be easily detected by Western-Blot. Again different concentrations of

DDM were used to solubilize the SecDF-SPA cell membranes. To obtain better results we have loaded

the maximum quantity of protein possible (in this case 17.5µg, all the samples were loaded with same

amount) and to avoid contamination between samples, we have loaded the samples with four empty

wells between. In BN-PAGE, proteins often appear as a smear and not as single bands as in SDS-

PAGE gels (Figure 29). Nonetheless, at low DDM concentrations the smears corresponding to

SecDF-SPA appeared higher in the gel, indicating that SecDF-SPA is part of a higher molecular

weight complex. In contrast, at higher DDM concentrations this complex is dissociated as SecDF-SPA

appeared lower in the gel (Figure 29).

Chapter 2.RESULTS

72

Figure 29. Analysis by Western blot of a Blue native PAGE 4-16% Bis-Tris Gel of SecDF-SPA in the

presence of different concentrations of DDM. The triangle on the top represents the increase of DDM

concentration, from 0.05% up to 2%. The red circles represent the loaded samples, which with the increase of the

DDM concentration decreases its position in the gel.

For the analysis of BN-PAGE by LC-MS/MS each gel lane was divided in ten small pieces,

in-gel trypsin digested to identify proteins present in each part of the lane (Figure 30). The results we

obtained from the LC-MS/MS analysis of these samples indicated that 1% DDM is the best

concentration to solubilize the membrane samples of B. subtilis, as it was the condition with a higher

number of proteins detected. We needed a membrane protein complex that can be used as a control in

our experiment, and, according to preliminary data obtained at our laboratory SppA and YteJ could

co-migrate together in BN-PAGE. In our experiment, in the presence of 1% DDM, the LC-MS/MS

results showed that the presence together of SppA and YteJ in the gel fraction number 4 (Figure 30).

Interestingly, YteJ was identified in the piece of gel in which the highest number of SppA peptides

were detected. At lower or higher DDM concentrations, YteJ was not detected.

Unfortunately, peptides belonging to SpoIIIJ were identified in almost all pieces of gel, with

an equal number of spectra in each fraction, indicating that SpoIIIJ might dissociate easily from a

possible membrane complex. Moreover, for an unknown reason, SecDF was not detected in any piece

of gel, in any DDM conditions (Figure 30). Altogether, these results reinforced the hypothesis that

SppA and YteJ form a stable complex, while, if SpoIIIJ and SecDF form a membrane protein

complex, it is unstable.

In conclusion, using SPA purification combined with BN-PAGE analysis, we were

unsuccessful to reveal any expected complexes of the Sec pathway. We focused on using native

73

condition to capture native complex; however, it might be necessary to use cross-linking agents to

study these complexes in B. subtilis.

As we have identified one complex with both techniques, the SppA/YteJ complex, the next

chapter will be entirely dedicated to the study of this membrane protein complex, that was supposed to

be related to the Sec pathway through the signal peptide peptidase function of SppA.

Figure 30. Number of peptides identified, by LC-MS/MS for each of the proteins purified in the presence

of 1% DDM. On the top of the figure, the gel lane represents how it is divided for in gel trypsin digestion and

posterior LC-MS/MS analysis. These graphs represents the number of peptides identified for SppA, YteJ,

SpoIIIJ and SecDF in function of each piece of gel.

Chapter 2.RESULTS

74

2.2 The SppA/YteJ membrane complex

The function of B. subtilis SppA has been studied, as described at the introduction (Signal

peptide peptidase A (SppA) section). However, yteJ which in an operon with sppA encodes for a

protein of unknown function and unknown structure. As a first step towards the understanding of this

unknown protein, we analysed (i) its genomic context using bioinformatics tools, such as BLAST

(Singh and Raghava 2016) and microbial genomic context viewer (Overmars et al. 2013), and (ii) its

putative structure using structure prediction softwares such as Protter (Figure S 1 of appendix chapter)

(Omasits et al. 2014). Using BLAST-P to align the sequence of the Bacillus subtilis sppA yteJ operon

with different bacterial genomes of different bacterial species, the results showed that yteJ is often

present following sppA. The sppA gene is not only present in B. subtilis but also in most of the bacteria

from the firmicutes phylum, including pathogenic bacteria as listeria monocytogenes (Figure 31).

Figure 31. Alignment of sppA and yteJ genes with other bacteria of firmicutes phylum. The red highlighted

box indicate the sppA yteJ operon. (Overmars et al. 2013).

2.2.1 The sppA yteJ operon is regulated by sigma factors σA and σW

Bacterial sigma factors interact with the RNA polymerase core and enable its binding to

specific promoter sequences, inducing the initiation of transcription (Britton et al. 2002). Sigma-A

(σA) is the major sigma factor required for expression of many essential housekeeping genes and the

majority of the RNAs necessary during bacterial growth (Britton et al. 2002).

The extracytoplasmic function (ECF) subfamily of sigma factors is a distinct group that allows

bacteria to adequately respond to stressful environmental conditions and that are involved in the

control of cell envelope-related functions, such as secretion, synthesis of exopolysaccharides or

macromolecules (Missiakas and Raina 1998). B. subtilis has seven members of this sigma factor

family, identified through bacterial genome sequencing. The sigma-W factor (σW), a member of this

subfamily, is normally induced in response to cell envelope stress caused by antibiotics, alkaline and

salt shocks (Zweers et al. 2012).

Chapter 2.RESULTS

75

In 1998, Huang and colleagues showed that the B. subtilis sppA and yteJ genes are members

of the σW regulon (Huang and Helmann 1998). Based on the data from Nicolas et al. (2012), the

expression of the B. subtilis sppA-yteJ operon seems to be dependent on σW but also possibly on σA,

the house keeping sigma factor. To test the relative importance of the σA and σW-dependent promoters

in the regulation of sppA yteJ expression, we constructed transcriptional fusions with these promoters

and the sfGFP (superfolder GFP). One construct possessed the sequence of the sppA yteJ σW-

dependent promoter, a fragment of 100 bp upstream the sppA gene and including the natural RBS

(Figure 32). A second construct possessed both σA and σW-dependent promoter sequences, a fragment

85 bp longer than the previous fragment, later denoted σAW. The promoter positions and lengths were

defined based on supplementary data provided in Nicolas et al. (2012).

Figure 32. Transcriptional fusions of the PσAW and PσW, promoters and sfGFP. PσW strain contain a fusion

of a fragment of 100bp upstream sppA and PσAW strain a fragment 185bp upstream sppA. Both constructions

harbour the sppA yteJ native RBS. These constructs were inserted at the ectopic amyE locus together with a

Spectinomycin resistance cassette.

These constructs were inserted ectopically at the amyE locus together with a spectinomycin

resistance cassette. The constructs were confirmed by PCR and sequencing. The final strains

harbouring either the σAW-dependent promoters or the single σW-dependent promoter were named

GHBs48 (BSB1 amyE::Paw-sfgfp) and GHBs 54 (BSB1 amyE::Pw-sfgfp), respectively. For the sake

of clarity, these strains were labelled PσAW and PσW, respectively. By following the fluorescence

intensity of the sfGFP, directly related to its expression, during growth we estimated the relative

quantitative contribution of each promoter in the sppA yteJ expression. As described above, the σW

factor is involved in cell-membrane and cell-wall stress, therefore we used two antimicrobial peptides

that perturb the cell membrane integrity, nisin and subtilin, to induce its activation. Nisin and subtilin,

two members of the class I of the lantibiotic family, insert themselves at the cell membrane by binding

to the lipid II, a peptidoglycan precursor. They arrest the Lipid II in the cell membrane, abrogating the

peptidoglycan formation (Cotter, Ross, and Hill 2012; Spieß et al. 2015). Consequently these peptides

induce the formation of pores in the cell membrane and affect the cell wall formation, which leads to

cell weakening and lysis (Cotter, Ross, and Hill 2012). Since the σW-dependent promoter is fused to

the sfgfp in both constructions, inducing its expression will be revealed by the sfGFP production level,

and therefore the contribution of the σA can be inferred. We tested a range of different concentrations

of these peptides (from 0 to 6 µg of nisin, and from 0 to 30% v/v of subtilin), and the results are shown

Chapter 2.RESULTS

76

in Figure 33 and Figure 34. The strains were grown in LB medium, supplemented with 100 µg.mL-1

spectinomycin, at 37°C with vigorous shaking in flasks. Overnight cultures were diluted 20-fold in

100 µL per well of pre-warmed fresh CH defined medium (pre-culture) into 96-well microtiter plates.

The CH pre-cultures were grown until an OD600nm of 0.3-0.4 and then diluted 400-fold into a pre-

warmed fresh medium. The antimicrobial peptides were added when the cell cultures reached early

exponential phase, around 0.2 OD600nm. OD600nm and fluorescence were recorded at time intervals of 10

minutes. The Figure 33 and Figure 34 display the PσAW and PσW strains growth curves and the

fluorescence/OD600nm ratio of sfGFP after induction (time point 0 min) with nisin or subtilin,

respectively. The background signal and the auto-fluorescence intensity were subtracted from all the

graphs.

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77

Figure 33. Nisin induced increasing expression of sfGFP, regulated by σA and σW sigma factors.

Transcriptional fusion of sfgfp with the full σA and σWdependent promoter sequence (PAW) or only the

σWdependent promoter sequence (PW) found upstream of sppA yteJ operon. The sfGFP expression was induced

with different concentrations of nisin, from 1µg to 6µg. Each graph line corresponds to a specific nisin

concentration, labelled on the left of the panel. The cell growth is shown on the left column of the panel while

the sfGFP fluorescence intensity /OD600nm is presented on the right. The dashed black line highlights the intensity

peak always found 60min after induction. represents the PσAW and PσW growth (left panel) and

fluorescence/OD600nm (right panel).

Chapter 2.RESULTS

78

At the top of Figure 33, the graphs labelled 0 µg corresponds to the controls, i.e. the PσAW and

PσW strains growing in LB without nisin. Both strains showed identical growth rates, while the

intensity of measured sfGFP fluorescence remained low. With 1 µg of nisin (second line panel of

Figure 33), although the three characteristic phases of the bacterial growth (exponential, transition and

stationary phase) can be observed, when compared with the controls 0 µg, the growths of both strains

are affected. Just after induction, a decrease in the cell growth was observed followed by a rapid

growth recovery. However, these cells do not reach the same optical density as the controls and as in

stationary phase, the OD of the nisin-treated cultures showed a slight decrease due to cell lysis. When

treated with 2 µg of nisin the strain growths were affected, as a drop in the OD600nm was measured just

after induction. Probably a part of the cells died, by cell lysis, and it took around 200 min to the

remaining living cells to produce enough biomass to be measurable by OD. The corresponding ratio of

sfGFP fluorescence intensity/OD600nm showed, 60 min (represented by a dashed line) after the

induction, a clear peak, meaning that the remaining living cells were producing higher amount of

sfGFP. This was interpreted as a boost of activity of the sigma-dependent promoters. For higher

concentrations of nisin, from 3 µg up to 6 µg, a longer lag phase was observed during which the

OD600nm was not measurable. Regardless of nisin concentrations, the levels of the fluorescence/OD

ratio increased during the first 60 min. The highest levels were recorded with 3 µg.ml-1 of nisin, while

the addition of 4 or 6 µg.ml-1 of nisin led to lower levels. For 2 and 3 µg.mL-1 of nisin, a peak of

fluorescence/OD ratio was observed - i.e. same duration for the increase and decrease of the signal,

while for higher concentrations (4 and 6 µg), a longer lag phase and even a plateau than what we

observed for lower nisin concentrations. Whatever the nisin concentration applied to the cells, the

fluorescence/OD600nm ratios followed the same patterns and levels in the two strains, PW and PAW.

Therefore, whether the sfgfp gene was under the control of a single promoter sequence, i.e. in the PW

strain, or the two adjacent promoters sequences found natively in the chromosome, i.e. in the PAW

strain, the results were identical. The exception was a slight increase of this ratio that was observed for

the highest concentrations of nisin (6 µg.ml-1), which indicated a very marginal contribution of A to

the overall control of the operon expression. We concluded that the main promoter responsible for the

genetic regulation of the sppA yteJ operon was the W-dependent promoter. It should be noted that the

fluorescence/OD ratio decreased to reach back the minimal levels after a period of time which

increased proportionally to the nisin concentration. Moreover, whatever the nisin concentration

considered, when the signal ratio was back to the minimum level, the cell growth concomitantly

resumed. As W is activated in response to nisin, one might suggest that the nisin concentrations

decreased below the activation threshold of W, and that these concentration levels were compatible

with cell division.

Chapter 2.RESULTS

79

Figure 34. Subtilin induced increasing expression of sfGFP, regulated by σA σW sigma factors. Transcriptional fusion of sfgfp.with the σA and σWdependent promoter sequences, native of the sppA yteJ operon.

The sfGFP expression was induced with different concentrations of of subtilin, from 5% up to 30%. Each graph

line corresponds to a specific subtilin concentration, labelled on the left of the panel. The cell growth is shown

on the left column of the panel while the sfGFP fluorescence intensity /OD600nm is presented on the right. The

dashed black line highlights the intensity peak always found 60min after induction. represents the PσAW

and PσW growth.

Chapter 2.RESULTS

80

Similar results were obtained when cells were treated with subtilin (Figure 34), an

antimicrobial peptide of the same family. At low concentration of subtilin (5%), we did not observe

any difference between the control and the treated strains (first and second lines of the Figure 34),

neither for the growth nor for the sfGFP production. When we used 10% of subtilin, the sfGFP

expression was induced, displaying a small peak in the fluorescence intensity. When increasing the

treatment to 15% of subtilin we reproduce the results observed with 2 µg of nisin. After 60 min

(represented by the dashed line) of subtilin treatment, a clear peak on the sfGFP fluorescence intensity

ratio was observed. As observed for nisin, treating the strains with high concentrations of subtilin,

20% and 30%, leads to an increase in the lag phase and a W-dependent response 60 min after

induction, with higher and wider peaks of fluorescence intensity. In the presence of 30% of subtilin,

cells did not grow. but a high fluorescence peak with a larger plateau, than the lower concentrations of

subtilin, was observed. In conclusion, we observed that regardless of the nisin or subtilin

concentrations, the major contributor for the regulation of the expression of the sppA yteJ operon was

σW.

2.2.2 Deletion of the sppA and yteJ genes

To better understand the role of the B. subtilis SppA/YteJ complex, we deleted these two

genes either separetly or together to study the effect of their deletion(s). The sppA, yteJ and both genes

were deleted from their native locus and replaced by a chloramphenicol resistance cassette (cat). The

expression of this cassette remained under the control of the sppA yteJ native promoter, as described at

the section 4.3.2 of the Material and Methods chapter. These constructs were confirmed by PCR and

sequencing. The final strains were named: GHBs 023 (BSB1 ∆yteJ::cat), GHBs 025 (BSB1

∆sppA::cat) and GHBs 028 (BSB1 ∆sppA yteJ::cat). For the sake of clarity, strains GHBs 023, 025

and 028 will be named ∆yteJ, ∆sppA and ∆sppA yteJ, respectively. The growth of the strains were

recorded by measuring, at regular intervals, the optical density at 600 nm (OD600nm). The Figure 35

shows the growth curves plotted for each strain. The wild type strain (BSB1) and the deletion mutants

presented the same growth rate and cell density at all growth stages. These results allowed us to

conclude that the deletion of the sppA and/or yteJ genes did not affect the growth of the mutant strains

in LB medium. This is in agreement with the result reported in Bolhuis et al (1999) for the deletion of

sppA.

Chapter 2.RESULTS

81

Figure 35. Growth of B.subtilis sppA and yteJ deletion mutant strains. The control strain (BSB1) culture was

grown overnight in LB and the mutant strains (∆yteJ, ∆sppA and ∆sppA yteJ) grown in LB supplemented with

5 μg.ml-1 chloramphenicol at 37°C, and the next day diluted 1:400 into a new medium with the same conditions.

We checked by microscopy whether cell morphology was affected. Cells were grown exactly

as previously described and samples were collected during the exponential (exp), transition (tran) and

stationary (stat) phases of the growth cycle and analysed by microscopy (Figure 36). Cells were

stained with the membrane dye FM4-64, allowing the observation of the cell membranes and septum.

The cell morphology of the ∆sppA yteJ strain was not affected, as shown in Figure 36. In exponential

phase, first line of Figure 36, comparing the cells of the BSB1 strain (on the left side of the panel)

with the ∆sppA yteJ mutant cells (right side of the panel), both appear to be normal B. subtilis rod-

shaped growing cells with 4-10 μm long (Yu et al. 2014), with a well-defined division septum and cell

poles. Identical cell morphology was observed in transition and stationary phases. Cell morphology of

each single deletion mutant strains were also checked by microscopy and as the double deletion

mutant no phenotypic differences were observed comparatively to the BSB1 strain. With these results

we concluded that the deletion of sppA, yteJ or sppA yteJ do not affect cell morphology.

Chapter 2.RESULTS

82

Figure 36. Cell morphology of the sppA yteJ deletion mutant. The strains, control strain (BSB1) and

∆sppA yteJ, were grown overnight in LB at 37°C, supplemented with 5μg/ml chloramphenicol when required,

and in the next day diluted 400-fold into fresh medium with the same conditions. Samples were collected during

the exponential (exp), transition (tran) and stationary (stat) phases for microscopy analysis. Phase contrast

microscopy pictures and corresponding red fluorescence microscopy (515/640nm) are shown. Membranes (false

coloured red) were stained with FM4-64. Scale bar is 10 micron.

2.2.3 Role of the SppA/YteJ complex in protein secretion

Bolhuis et al., (1999) showed that the deletion of sppA affected the maturation of a highly

secreted protein, AmyQ. AmyQ, a secreted protein through the Sec pathway, is maintained in an

unfolded pre-protein form in the cytoplasm and reaches its folded mature form after being translocated

through the cell membrane and released to the extracellular medium. Bolhuis et al., (1999) observed

that the sppA deletion caused a delay in the processing of the AmyQ mature form which led to an

accumulation of the pre-AmyQ, which resulted in a mild decrease in the secretion of AmyQ. These

results suggested that the SppA activity was important in the efficient processing of the pre-secreted

proteins, supposedly by degrading the signal peptides from secreted proteins (Bolhuis et al. 1999).

While the role of SppA in protein secretion was documented, we wondered what was the role of YteJ,

and also what was the role of the SppA/YteJ complex in this cellular process. To address these issues,

Chapter 2.RESULTS

83

we checked the secretion levels of a highly secreted protein, the maltogenic amylase (AmyM). Dr.

Van Rij, from DSM (Delft, The Netherlands), provided us with a multi copy plasmid containing the

gene encoding AmyM (pCS73) and the corresponding empty plasmid (pCS74), all containing a

neomycin (neo) resistance cassette. Each one of these plasmids was transformed into the B. subtilis

sppA, yteJ or sppA yteJ deletion strains and the BSB1 wild type strain, giving rise to eight new

strains (listed in Table S5 in the appendix section) from GHBs029 to GHBs040. AmyM protein has a

molecular weight of 75.2 kDa and can be followed by SDS-PAGE.

To analyse the extracellular fraction, the cells were harvested in exponential phase of growth

and centrifuged. The obtained cell-free supernatant was supplemented with TCA to precipitate the

extracellular secreted proteins that were then analysed by SDS-PAGE. Before running the gel, the

concentration of the protein samples were measured by the Bradford method and equally loaded. The

results are shown in Figure 37. The two first lanes correspond to the control samples, the BSB1 wild-

type strain and BSB1 with pCS74, respectively. These two strains do not contain the amyM gene. The

sample BSB1 containing pCS74 empty plasmid was used as a control to check if the presence of a

multicopy plasmid affects the normal protein content. Compared to the BSB1 sample without plasmid,

first lane, the presence of the empty plasmid do not affect the protein content, as both lanes showed the

same protein pattern. The remaining four lanes on this gel correspond to the extracellular protein

samples of the strains expressing AmyM, thus containing the pCS73 plasmid, including the respective

control BSB1 and deletion mutant strains. All these samples showed a band around 75 kDa that

corresponds to the molecular weight of AmyM.

All three deleted strains showed the same intensity for the AmyM band on the gel. This

indicated that the deletion of sppA, yteJ or the operon sppA yteJ had no impact on the overall amount

of the hyper-secreted AmyM protein.

The delay in the processing of the AmyQ mature form observed by Bolhuis et al., (1999) was

a mild effect and the results were obtained through pulse-chase protein labelling. These facts can

explain why it is not possible to see a difference on the final amount of the mature form in our

experiments. To further study the premature and mature form of the secreted protein and analyse the

differences between the amount of secreted protein in the wild-type and mutant strains, we needed to

use specific antibodies for this protein. The production of AmyM antibodies, from the protein

purification up to rabbit immunization, was part of a secondment project developed in collaboration

with one of our industrial partners of the ProteinFactory ITN, UCB-Celltech company (Slough, UK),

including an internship of six weeks on UCB premises. The details of this work are described in the

Appendix 6.2.

Chapter 2.RESULTS

84

Figure 37. Analysis of AmyM extracellular amount by SDS PAGE. pCS74 empty plasmid; pCS73 plasmid

encoding amyM gene. The BSB1 wild type strain, BSB1 with pCS74 empty plasmid and with the pCS73 AmyM

plasmid were used as controls. The black arrow points for the correct band of AmyM.

With the serum of antibodies against AmyM we analysed the extracellular fraction, obtained

as previously explained, by Western Blot (WB). To do the WB we ran the samples in a 10%SDS-

PAGE gel exactly as described for the previous experiment. In the first two lanes, corresponding to the

controls, we cannot (as expected) observe a band corresponding to AmyM, since these strains do not

produce AmyM. The remaining samples, expressing AmyM, showed a band around 75KDa

corresponding to the molecular weight of AmyM and some lower bands corresponding to nonspecific

antibodies reaction or possible AmyM degradation. However, once more, this technique did not show

significant or measurable differences in the amount of secreted AmyM, between the strains. We also

tried to use a second reporter protein, the highly secreted endo-1,4-beta-xylanase (XynA). The xynA

gene was also provided by Dr. Van Rij, on a multi copy plasmid, and we proceeded to the same

experiment as just described above for AmyM. Unfortunately, it was not possible to detect XynA on

SDS-PAGE , maybe due to low expression and concentration of this secreted protein in the medium.

Chapter 2.RESULTS

85

Figure 38. Analysis of AmyM extracellular amount by Western Blot. This WB was performed with

antibodies against AmyM. pCS74 empty plasmid; pCS73 plasmid encoding amyM gene. The BSB1 wild type

strain, BSB1 with pCS74 empty plasmid and with the pCS73 AmyM plasmid were used as controls. The black

arrow points for the correct band of AmyM.

2.2.3.1 Effect of the sppA, yteJ or sppA yteJ overexpression in the secretion

Since we could not observe a difference in secretion caused by the deletion of the sppA and

yteJ genes, we wondered what would be the effect of their overexpression. Therefore, we constructed

overexpressing strains by re-inserting, in the WT BSB1 background, each gene or the full operon in

the amyE ectopic locus under the control of the hyperspank promoter (Phs), an IPTG inducible

promoter (as described in the material and methods section 4.3.3). These constructs were confirmed by

PCR and sequencing. The final strains are GHBs 059 (BSB1 amyE::Phs yteJ /spec), GHBs 066 (BSB1

amyE::Phs sppA /spec) and GHBs 071 (BSB1 amyE::Phs sppA yteJ /spec). For the sake of clarity, the

strains GHBs 059, 066 and 071 will be named BSB1 PhsyteJ, BSB1 PhssppA and BSB1 PhssppA yteJ,

respectively. To address the effect of the overexpression of SppA, YteJ and of the SppA/YteJ complex

in protein secretion we transformed these strains with either the multi copy plasmids containing the

gene encoding AmyM (pCS73) or an empty plasmid (pCS74) as a control, as we proceeded before

with the deletion mutants. This way, we could followed by SDS-PAGE the levels of AmyM secreted

into the extracellular medium. It gave rise to nine new strains (listed in Table S5 in the appendix

section) from GHBs073 up to GHBs081.

The overexpressing strains, listed above, were grown in LB at 37°C, supplemented with the

correct antibiotics when needed, and the cells were harvested in exponential phase and centrifuged.

The extracellular cell-free medium was TCA precipitated and analysed by 10% SDS-PAGE gels.

Before loading the samples onto the gel, the concentrations in total protein were measured by the

Chapter 2.RESULTS

86

Bradford method and, then, loaded with the same concentration of 0.5 µg.µl-1. These results are

shown at Figure 39.

Figure 39. SDS PAGE analysis of AmyM extracellular amount produced by B. subtilis WT and

overexpression mutant strains. This picture corresponds to a 10% SDS-PAGE gel, coomassie staining. The

BSB1 wild type strain without and with the pCS73 AmyM plasmid were used as controls. BSB1 PhsyteJ, BSB1

PhssppA and BSB1 Phs sppA yteJ stands for B. subtilis yteJ, sppA and sppA yteJ overexpression mutant strains,

respectively. pCS73 multi copy plasmid encoding amyM gene. The black arrows highlight the 85 kDa band of

the protein standard and the band corresponding AmyM (75.2 kDa).

The second lane of the SDS-PAGE shown on Figure 39 corresponds to the extracellular

medium sample of the BSB1 control strain, without the pCS73 (AmyM) plasmid. As this strain do not

contain the amyM gene, AmyM is not produced, so the band around 75 kDa corresponding to AmyM

is absent. In contrast, BSB1 with the pCS73 plasmid, on the third lane, shows a band with the expected

size for AmyM, confirming that AmyM is produced. However, in the fourth lane, we can observe that

when sppA is overexpressed the amount of AmyM in the extracellular medium is significantly

reduced, as the 75 kDa band, corresponding to AmyM, present in this sample is fainter than the band

shown on the third lane control. Interestingly, comparing to the control sample, when yteJ or sppA yteJ

are overexpressed the amount of AmyM present in the extracellular medium increases. The last two

lanes of the gel (Figure 39), corresponding to the yteJ or sppA yteJ overexpressing samples

respectively, show a AmyM band more intense than the AmyM band present in the control of the

second lane. With these results, we can conclude that when the amount of SppA increases in the cell

Chapter 2.RESULTS

87

the amount of secreted AmyM decreases. As SppA is a membrane protease, it might degrade directly

the AmyM produced or it could be an indirect effect. When YteJ is overexpressed, AmyM levels seem

to be higher than those observed with BSB1 strain, as if AmyM was less degraded. Furthermore, when

SppA is co-overexpressed with YteJ, the levels of AmyM are higher than BSB1. If SppA is able to

degrade AmyM directly – not yet proven – one can hypothesize that the SppA protease activity might

be inhibited or regulated by YteJ.

2.2.4 The deletion of sppA yteJ affects the long term survival

The ability to replicate in a specific environment is named bacterial fitness. The fitness of a

bacteria can be tested by analysing the capacity of survival of two strains, one antibiotic sensitive

strain and one antibiotic resistant strain, under the same conditions for a predefined time. Furthermore,

at the end of the predefined time, the number of bacteria is counted and the one which has produced a

higher number is by definition "Fitter".

In order to characterise the ∆yteJ, ∆sppA and ∆sppA yteJ deletion mutant strains we studied

their capacity of survival by performing some fitness tests. First the strains were grown in liquid rich

medium (LB), at 37°C for 48h. At different time points (after 18h, 24h and 48h), samples of the

cultures were plated out onto LB agar plates. The number of colonies were counted after the plates

have been incubated for 18h at 37°C. As shown in Figure 40, after 18h of growth, the number of

colonies counted is just slightly affected by the deletion of sppA, yteJ and the full operon, as compared

with the number of WT colonies. The lowest number of colonies was counted for the ∆sppA strain,

showing around 200 CFU/ml, which is 1.5-times less than the wild type. After 24h of growth in the

same medium, the capacity of survival of all strains was affected, because the number of colonies was

lower than the numbers after 18h of growth, even for the WT control strain. However, the capacity of

survival of the deletion mutants was even more affected, as in the absence of these genes the number

of surviving colonies is lower than the WT. The ∆sppA strain was the more affected, showing the

lowest value of colonies counted, around 150 CFU/ml lower than the wild type control.

After of 48h of growth, as shown on Figure 40, although all the strains showed a lower

number of surviving colonies, the mutant strains have a lower capacity of survival. The ∆sppA yteJ

and ∆sppA strains are both the less fit, as both have the lowest number of colonies counted. The yteJ

strain was the least affected strain compared to the other mutant strains. The results of these tests

showed that in LB medium the survival of the deletion strains is affected comparing to the wild type

strain. We concluded that the deletion of sppA was the most deleterious one for survival, indicating

that SppA function provides an advantage for long term survival. The same conclusion applied to the

deletion of sppA yteJ operon. However, as the deletion of yteJ was the less deleterious, we concluded

that YteJ function was not important for survival.

Chapter 2.RESULTS

88

Figure 40. Capacity of survival of BSB1 and ∆yteJ, ∆sppA and ∆sppA yteJ deletion mutants. The strains

were grown in rich liquid media (LB), at 37°C for 48h. After 18h, 24h and 48h, samples of the cultures were

collected and plated out onto LB agar plates. The number of colonies were counted after the plates have been

incubated for 18h at 37°C.

The survival ability of the ∆sppA yteJ mutant was as much affected as for the ∆sppA mutant

strain. As our main objective was to understand the function of the SppA/YteJ complex we performed

further fitness tests with only BSB1 and ∆sppA yteJ strains. We wondered whether the composition of

the media may affect the long-term survival of the strains. New fitness tests in LB rich media (Figure

41A) and in M9 minimal medium (M9 supplemented with glucose) were realised (Figure 41B). After

24h and 48h, samples were collected and plated on LB agar plates, and colonies were counted after

18h of growth at 37°C.

Chapter 2.RESULTS

89

Figure 41. Long-term survival of BSB1 and ∆sppA yteJ deletion mutant, in LB and M9 liquid media. (A)

Strains grown in LB, a rich liquid media; (B) strains grown in M9 minimal medium. Strains were grown at 37°C

for 48h. After 24h and 48h, samples of the cultures were collected and plated out onto LB agar plates. The

number of colonies were counted after the plates have been incubated for 18h at 37°C.

Figure 41A shows the long-term survival of BSB1 and the ∆sppA yteJ strains. The deletion of

the sppA yteJ operon clearly affected the long-term survival of the cells. Figure 41B shows the long-

term survival of the same strains in M9 minimal medium. After 24h in the M9 medium, the number of

CFU/ml for BSB1 was the same as when cells were grown in LB; however the number of CFU/ml for

∆sppA yteJ was already half of that observed in LB. After 48h, the CFU/ml number for BSB1 was half

of that observed in LB indicating that the M9 medium is not as good as LB for long-term survival,

even for the wild-type cells. After 48h in the M9 medium, the ∆sppA yteJ cells showed the lowest

number of surviving cells. We can conclude that the medium composition and richness affect the long-

term survival of B. subtilis cells and that this effect is more pronounced for the ∆sppA yteJ strain.

Bolhuis et al. in 1999 showed that the deletion of B.subtilis sppA affected protein secretion.

Taking in account this fact, and what we observed in our results, we wanted to test if the BSB1 strain

secreted a protein whose function or quantity was altered in the ∆sppA yteJ strain. We performed

complementation assays for which the WT and ∆sppA yteJ strain were grown together in LB medium,

at 37°C. Using the same procedure as described above, separated cultures of BSB1 and ∆sppA yteJ

strains, as well as a mix of 50% BSB1 and 50% ∆sppA yteJ cells were grown in the same conditions.

After 24 and 48h of growth in the LB medium, cells were diluted and plated on LB agar medium.

However the mixed culture was plated on a second plate of LB agar supplemented with

chloramphenicol (Cm) to count the Cm-resistant cells, i.e. the ∆sppA yteJ cells. The results are

presented in

Figure 42.

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Figure 42. Complementation assays in LB with BSB1 and ∆sppA yteJ deletion mutant strains. (A) results

obtained after 24h of growth; (B) results obtained after 48h of growth; BSB1 strain was plated in LB agar;

∆sppA yteJ mutant strain was plated into LB agar supplemented with Cm 5µg/ml; Comp- stands for

complementation mixed sample. A mixture of 50% BSB1 and 50% ∆sppA yteJ composes this sample. Comp1

shows the number of colonies counted when the Comp sample is plated into LB agar and Comp2 LB agar

supplemented with Cm 5 µg/ml.

For both 24 and 48 hours of growth (

Figure 42) we observed that the ability to survive of the ∆sppA yteJ strain is lower than that of

the WT strain. For the complementation experiment with the inoculation using an equal number of

BSB1 and ∆sppA yteJ cells, after 24h, the total number of cells was approximately twice that of the

∆sppA yteJ cells. This indicated that during the first 24h, cells of both strains grew equally well.

However after 48h, the ∆sppA yteJ cells represented only 15% of the total cells. This experiment

indicated that the defect of long-term survival of the ∆sppA yteJ cells was not due to the lack of a

specific secreted protein or another secreted component in the medium, as BSB1 cells did not

complement it.

The general conclusion is that the function of SppA is important for the long-term survival of

B. subtilis cells and, that, even if the protease active sites of SppA are located outside the cell, SppA

activity cannot help the survival of other cells around, but this activity seems important for the cell

itself.

2.2.5 The deletion of sppA and yteJ affects the resistance to lantibiotics

In 2013, Kingston and his colleagues demonstrated that the deletion of B. subtilis SppA leads

to a strain sensitivity to nisin, an antimicrobial peptide (Kingston, Liao, and Helmann 2013a). To

investigate the possible involvement of the SppA/YteJ complex in the resistance to antimicrobial

peptides we tested the resistance capacity of deletion ∆sppA and ∆yteJ deletion mutant strains to nisin

and subtilin. Nisin and subtilin are two antimicrobial peptides of the same lantibiotic family (Spieß et

al. 2015). This experience was performed in 96-well plates, where the strains grew over 16 hours in

the presence of nisin or subtilin, optical density at 600nm (OD600nm) was measured at regular intervals.

The BSB1 strain was used as a control strain. Overnight cultures of BSB1 and ∆sppA, ∆yteJ and

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91

∆sppA yteJ strains were grown at 37 ᵒC, in LB and LB supplemented with Cm 5 µg/ml respectively,

with aeration and vigorous shaking in flasks. The overnight cultures were diluted 20-fold into 100µl

(into the 96 well plate) of fresh pre-warmed LB medium and grown until beginning of exponential

phase. Then the cells were re-diluted 400-fold into a new plate, containing different concentrations of

nisin or subtilin. The Figure 43 shows the results obtained with nisin.


concentrations of Nisin. The BSB1, ∆sppA and ∆yteJ strains grown in LB in a 96-well plate, where the OD600

was measured at specific time points. The different graphs represent the growth of the strains in the presence of

different concentrations (2 µg, 3 µg, 4 µg, 6 µg and 8 µg/ml) of nisin, labelled on the left of the graph.

In Figure 43, BSB1, ∆sppA and ∆yteJ strains showed identical growth rates when grown in

LB without nisin. With 3µg/ml of nisin (graph middle left), although the three characteristic phases of

the bacterial growth (exponential, transition and stationary phase), the growth of the strains was

affected. The strains took around 100 extra minutes to resume growth showing a longer lag phase than

in the control conditions. We defined the time necessary to resume growth as the time needed for the

strain to reach an OD600 of 0.1. In the presence of 3µg/ml of nisin, the deletion of yteJ led to a slightly

increased resistance to nisin, as it took around 20 minutes less than the BSB1 strain to reach an OD600

of 0.1. On the contrary the deletion of sppA turn the cells more susceptible to nisin, as it took around

20 minutes more than the BSB1 strain to reach an OD600 of 0.1. When the concentration of nisin was

increased to 4 µg/ml in the medium, the same results were observed but with an exacerbate difference

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92

between the strains. When treated with 6µg/ml of nisin the BSB1 strain showed a lag phase of 250

minutes longer than in the control condition without nisin. In this condition, the ∆yteJ strain showed

the same lag phase and growth rate as the BSB1 strain, while the ∆sppA strain took slightly longer to

resume growth. The ∆sppA strain is therefore more susceptible to nisin than the two other strains. In

the presence of 8µg/ml of nisin, the strains did not grow.

To simplify we did not represent the ∆sppA yteJ results, as this mutant strain behaved exactly

as the yteJ mutant strain. With these results we concluded that the deletion of sppA affects de

resistance to nisin. However the deletion of yteJ or the full operon, yteJ sppA, showed non

reproducible results with either an intermediate phenotype or a nisin resistance higher than that of the

BSB1 strain.

Although the trend for the nisin resistance of the BSB1 strain compared to the deletion strains

were reproducible, the effect of a specific nisin concentration could vary from one experiment to

another. The stability of nisin might explain these differences. Therefore we decided to test another

lantibiotic of the same class with a highly similar ring structure, subtilin. (Spieß et al. 2015). The

experiments were performed as explained before for nisin. As shown in Figure 44, the results

obtained with subtilin were similar to that observed with nisin.

Figure 44. Growth of the B. subtilis sppA and yteJ mutant strains in the presence of different

concentrations of subtilin. The BSB1, ∆sppA and ∆yteJ strains were grown in LB in a 96-well plate, and the

OD600 was measured at specific time points. The different graphs represent the growth of the strains in the

presence of different concentrations (20%, 25%, 30%, 34% and 38%) of subtilin, labelled on the left of the

graph.

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93

With 20% of subtilin (graph top right, Figure 44), although we observed the three

characteristic phases of the bacterial growth (exponential, transition and stationary phase), when

compared with the control conditions 0% (graph top left), the growth of the strains was altered. The

cells took slightly longer to resume growth, did not reach the same optical density in stationary phase

as in the control conditions. The growth of the two deletion strains showed a small delay compared to

the BSB1 strain. When treated with 25% of subtilin, the BSB1 and ∆yteJ mutant strains had identical

growth rates, and similar 'lag phase' of around 200 min. The ∆sppA strain showed an even longer lag

phase, around 100 extra minutes to resume growth, indicating a higher susceptibilty to subtilin.

Increasing the subtilin to 30% led to a striking difference between the growths of the three strains.

Comparing BSB1 with ∆yteJ, we can see that the deletion of yteJ led to a decrease in the resistance to

subtilin, as it took around 200 minutes more than the BSB1 strain to reach an OD600 of 0.1. The

deletion of sppA turn the strain even more susceptible, as it took 300 minutes more than the BSB1

strain to reach an OD600 of 0.1. The deletion of yteJ showed an intermediate phenotype, between the

BSB1 and ∆sppA strains. When the concentration of subtilin was increased to 34%, the BSB1 strain

was able to resume growth after 300 min while it took 600 min for the ∆sppA strain and while the

∆yteJ strain showed almost no growth. Altogether these results led us to conclude that the deletion of

either yteJ or sppA turned the cells more susceptible to subtilin. In the presence of 38% subtilin, the

strains did not grow. As stated before for the nisin results, with subtilin we did not show the

∆sppA yteJ results, as this strain had the same behaviour as the yteJ strain. With these results,

comparable to the nisin results, we can conclude that the deletion of sppA affects de resistance to

subtilin. However, depending on the concentration of subtilin, the deletion of yteJ or of the full operon

showed again non reproducible results (as for nisin) with either an intermediate phenotype or a subtilin

resistance higher than that of the BSB1 strain.

Figure 45 illustrates the effect of increasing subtilin concentrations on the growth rate. It

revealed that the growth rates were strongly affected when the concentration of subtilin was higher

than 30%. As the results obtained with subtilin were more reproducible than the experiments done

with nisin, and as a 30% subtilin concentration showed significant differences in the lag phase

duration of the three strains without affecting too much the growth rates (and so allowing to

discriminate strains based on their subtilin resistance capacity), we selected this concentration to setup

the lantibiotic resistance test used later in this work.

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94

Figure 45. The growth rates of BSB1, ∆sppA and ∆yteJ in the presence of different concentrations of

subtilin. The BSB1, ∆sppA and ∆yteJ strains were grown in LB in a 96-well plate, and the OD600 was measured

at specific time points. The different columns of the graph represent the growth rates of the strains in the

presence of different concentrations (20%, 25%, 30%, 35% and 40%) of subtilin.

2.2.6 The deletions of sppA and yteJ do not affect the resistance to

vancomycin, erythromycin or protect against lysozyme

Our results showed that the SppA/YteJ complex was involved, mainly through SppA, in the

resistance to lantibiotics, polycyclic antimicrobial peptides that are capable of inserting themselves

into the cellular membrane. To check whether the SppA/YteJ complex had a role in the resistance to

other classes of antibiotics we repeated the previous experiments with two other classes of antibiotics.

First, we chose to test an antibiotic of peptidic nature, ancomycin, a glycopeptide antibiotic which acts

by inhibiting the cell wall synthesis of Gram-positive bacteria. The experiment was performed as

described in the previous section 2.2.5, testing a range (from 0 to 1 µg/ml) of vancomycin

concentrations (Adimpong et al. 2012).

In Figure 46, the first graph (top left) of the panel corresponds to the controls, i.e. the BSB1

and ∆sppA and ∆yteJ mutant strains grown in LB without vancomycin. The three strains showed

identical growth rates. When treated with 0.1 µg of vancomycin (graph top right), the growth was not

altered, meaning that this concentration it is not high enough to induce a response. When 0.2 µg/ml of

vancomycin was used (graph middle left), the wild-type strain showed a long lag phase; the ∆sppA and

∆yteJ mutant strains did not grow. We observed similar results for higher concentrations of

vancomycin (0.5 and 1 µg/ml). Comparing these results with the results obtained for nisin and subtilin,

we can conclude that the deletion of sppA or yteJ does not affects the resistance to vancomycin. To

fully conclude on vancomycin, a range of concentrations between 0.1 to 0.2 µg/ml should be tested.

Chapter 2.RESULTS

95


concentrations of vancomycin. The BSB1, ∆sppA and ∆yteJ deletion strains grown in LB in a 96-well plate.

The OD600 was measured at specific time points. The different graphs represent the growth of the strains in the

presence of different concentrations (0µg, 0.1µg, 0.2µg, 0.5µg and 1µg) of vancomycin, labelled on the left of

the graph. Colours, on the right side of the picture, indicate the wild type or mutated strains.

To test a different class of antibiotic not related to cell wall or membrane synthesis or

integrity, we chose erythromycin. Erythromycin is part of the macrolide family of antibiotics, which

decreases the bacterial protein production. The experiment was performed as described earlier, testing

a range (from 0-1 µg/ml) of different concentrations of erythromycin, taking into account the minimal

inhibitory concentration (MIC) of this antibiotic against B. subtilis, 4 µg/ml (Adimpong et al. 2012).

The results obtained with erythromycin (Figure 47) are similar to that observed with vancomycin. In

the absence of erythromycin, the BSB1, ∆sppA and ∆yteJ strains showed identical growth rates.

However, when we started to treat with 0.1 µg of erythromycin (Figure 47, graph top right) strains did

not grow, meaning that at this concentration the antibiotic have a bactericidal effect. All the

concentrations used of erythromycin showed the same results. Furthermore, for all concentrations of

erythromycin used so far, the WT strain started to grow after 600 min, reaching an OD600 of 0.1.

Comparing these results with the results obtained with the lantibiotics, we concluded that the deletion

of sppA or yteJ do not affects de resistance to erythromycin.

Chapter 2.RESULTS

96


concentrations of erythromycin. The BSB1, ∆sppA and ∆yteJ deletion strains grown in LB in a 96-well plate.

The OD600 was measured at specific time points. The different graphs represent the growth of the strains in the

presence of different concentrations (0 µg, 0.1 µg, 0.2 µg, 0.5 µg and 1 µg) of erythromycin, labelled on the left

of the graph. Colours, on the right side of the picture, indicate the wild type or mutated strains.

SppA is a serine protease which is, according to Nam et al., 2012, capable of digesting fully

folded proteins. Therefore, we decided to test if the SppA/YteJ complex could be involved in a

mechanism that protects the cell against the activity of lysozyme, glycoside hydrolase that is used as

an antimicrobial enzyme due to its capacity to digest peptidoglycan, the major component of the

Gram-positive bacterial cell wall. The experiment was performed as previously described, testing

different concentrations of lysozyme, from 0 to 250 µg/ml. The lysozyme results are shown in Figure

48. Whatever the concentrations of lysozyme added in the medium, no marked differences were

observed between the BSB1 and the ∆sppA and ∆yteJ strains. (At 150 µg/ml lysozyme, the ∆yteJ

strain took slightly longer to reach OD600 0.1 compared to the BSB1 and ∆sppA strain. As the results

obtained for vancomycin and erythromycin, it seems that SppA/YteJ complex does not protect the

cells against the lysozyme action.

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Figure 48. The B. subtilis sppA and yteJ deletion mutant strains treated with different concentrations of

lysozyme. The BSB1, ∆sppA and ∆yteJ deletion strains grown in LB in a 96-well plate. The OD600 was

measured at specific time points. The different graphs represent the growth of the strains in the presence of

different concentrations (0 µg, 50 µg, 100 µg, 150 µg, 200 µg and 250 µg) of lysozyme, labelled on the left of

the graph. Colours, on the right side of the picture, indicate the wild type or mutated strains.

As a general conclusion about the role of SppA, we have confirmed that sppA is important in

the lantibiotic resistance mechanism; however the protecting effect of SppA against lantibiotics can

not be extended to any other antibiotic molecules that we have tested. The vancomycin peptide

structure (a tricyclic glycopeptide) might prevent SppA to degrade it; and SppA protease activity

seems to be unable to degrade the fully folded lysozyme. At this stage of the study, the role of YteJ in

the complex is more difficult to understand as, depending on the concentrations of lantibiotic used, the

positive or negative effect of the deletion of yteJ could be observed.

2.2.7 Overexpression of sppA results in the recovery of the BSB1 phenotype

in the presence of subtilin.

To check if the higher susceptibility to lantibiotics was specifically due to the deletion of sppA

and yteJ we constructed strains overexpressing these two genes, either individually or as an operon.

The overexpressing strains were constructed by re-inserting, into the mutant genetic background, each

gene or the full operon at the amyE locus under the control of the hyperspank promoter, an IPTG

inducible promoter. The final strains were named as GHBs 061 (BSB1 ∆yteJ::cat amyE::Phs yteJ

/spec), GHBs 067 (BSB1 ∆sppA::cat amyE:: Phs sppA /spec) and GHBs 072 (BSB1 ∆sppA yteJ::cat

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98

amyE::Phs sppA yteJ /spec). For the sake of clarity, strains GHBs 061, 067 and 072 will be named

PhsyteJ, PhssppA and PhssppA yteJ, respectively.

Cells were grown in 96 well plates, over 16 hours in the presence of 30% subtilin and different

concentrations of IPTG, and the optical density at 600 nm (OD600nm) was measured at regular intervals.

The BSB1 strain was used as the control strain. Overnight cultures of the BSB1 and PhsyteJ, PhssppA

and Phs sppA yteJ deletion mutant strains were grown at 37ᵒC with aeration and vigorous shaking in

flasks. BSB1 in LB and the overexpressing mutants in LB supplemented with 5 µg/ml Cm and

100 µg/ml Spec. The overnight cultures were diluted 20-fold into 100 µl (in the 96 well plate) of fresh

pre-warmed LB medium and grown until the beginning of the exponential phase. Then the cells were

re-diluted 400-fold into a new plate in the following conditions: LB as control, LB supplemented with

30% subtilin and LB supplemented with 30% subtilin and different concentrations of IPTG.

The Figure 49 shows the results for the overexpression of sppA and yteJ. In LB (0% subtilin

0 µM IPTG), the BSB1, PhsyteJ and PhssppA strains showed identical growth rates. In the presence of

30% subtilin and 0 µM IPTG, the PhssppA (no SppA is produced) showed a longer lag phase, around

100 extra minutes, compared to BSB1 and PhsyteJ. When 10 µM of IPTG was added, inducing the

overexpression of sppA and yteJ, we could observe that the lag phase of strain PhssppA decreased.

Increasing the concentrations of IPTG to 25 and 50 µM induced an even shorter lag phase for PhssppA

strain. The PhssppA resistance to 30 % subtilin was almost recovered at 50µM ITPG. With 50µM of

IPTG, the PhsyteJ showed a very slight increase in the resistance to 30% subtilin, as compared to the

BSB1 strain. However, when induced with 100 µM, the three strains first grew similarly but then the

high overexpression of sppA rapidly led to cell lysis in the stationary phase.

With these results, we can conclude that the deletion of sppA affects the resistance to subtilin

directly through the enzymatic activity of SppA, as the complementation with sppA overexpression

allowed the recovery of the subtilin resistance. However, a new phenomenon appeared with the early

cell lysis observed when 100 µM IPTG were used to induce sppA.

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99

Figure 49. The B. subtilis sppA and yteJ overexpressing strains. The BSB1, PhssppA and PhsyteJ

overexpression strains grown in LB in a 96-well plate, and the OD600 was measured on specific time points. The

different graphs represent the growth of the strains in different conditions: top left strains growing in LB; top

right- LB with 30% subtilin; remaining graphs: growths of the strains in 30% subtilin and different

concentrations (10 µM, 25 µM, 50 µM and 100 µM) of IPTG. The conditions are labelled on the top of each

graph.

We also tested the overexpression of the operon, PhssppA yteJ (

Figure 50). From 10 up to 100 µM IPTG, the subtilin resistance recovery of the PhssppA yteJ

strain was complete with the lowest concentration of IPTG (10 µM). However, the most remarkable

result was the fact that in the presence of IPTG the overexpression of the sppA yteJ operon showed a

shorter lag phase than the BSB1 strain. The PhssppA yteJ cells seemed to recover quicker from subtilin

action than the WT strain. These results reinforced the idea that YteJ may have a role in the regulation

of SppA, as when they were overexpressed together, the cells were more efficient to recover from

subtilin action. Moreover, the results brought another hint in the relation between the two proteins. As

the premature cell lysis, that was clearly observed when SppA was overexpressed alone, was not

observed when YteJ was co-expressed with SppA, we hypothesized that YteJ might negatively

regulate the activity of SppA, preventing its deleterious action.

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Figure 50. The B. subtilis sppA yteJ overexpressing strain. The BSB1 and PhssppA yteJ strains were grown in

LB in a 96-well plate, and the OD600 was measured at specific time points. The different graphs represent the

growth of the strains in different conditions: top left in LB; top right- 30% subtilin; remaining graphs: growth of

the strains in 30% subtilin and different concentrations (10 µM, 25 µM, 50 µM and 100 µM) of IPTG. The

conditions are labelled on the top of each graph.

2.2.8 The overexpression of sppA results in a change of the cell

morphology.

The overexpression of sppA led to premature cell lysis. As the effect was drastic, we checked

by microscopy if the overexpression of sppA, yteJ or sppA yteJ would affect the morphology of the

cells. The cells were grown overnight at 37 ᵒC with aeration and vigorous shaking in flasks, BSB1 was

inoculated into fresh LB media and the overexpression mutants were inoculated into LB medium

supplemented with 100 μg/ml Spec and 5 µg/ml Cm. Overnight cultures were diluted 400-fold into

fresh pre-warmed LB media without any supplements. Biomass (OD600) was measured at regular

intervals and samples were collected during the early stationary (stat) phase of the growth cycle and

observed by microscopy. The early stationary phase was chosen as, when induced with 100 µM IPTG,

at this growth stage it is possible to observed a growth defect for PhssppA cells (see Figure 49). Cells

were stained with the membrane dye FM4-64, allowing the observation of the cell membranes and

septum, and with DAPI, allowing the observation of chromosomal segregation. The cells of the BSB1

strain (Figure 51-first line on the panel) appeared to be normal B. subtilis rod-shaped growing cells, 4-

10 μm long (Yu et al. 2014), well-defined division septa, cell poles and well segregated chromosomes.

The overexpression of sppA affected cell morphology (second line Figure 51). The PhssppA cells were

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101

longer than the BSB1 cells, with septal formation problems and an abnormal chromosomal

segregation, as the cells did not divide equally and the chromosomal DNA was not well condensed in

the middle of the cell. On other hand, the PhsyteJ and PhssppA yteJ cells appeared as normal B. subtilis

rod-shaped cells with a well-defined division septa, cell poles and well segregated chromosomes. We

concluded that the presence of a high amount of SppA leads to a morphological and division defect.

However, when overexpressed together with YteJ, it seems that YteJ somehow counteracted the

morphological defect caused by the overexpression of SppA.

The observed phenotype, “elongated/early lysing cells” of the PhssppA strain, could be the

result of an membrane overloading by the overexpressed membrane protein SppA itself or by its

protease activity. To discriminate between these two hypotheses, we constructed an inactive form of

SppA and inserted it at the amyE locus under the control of the hyperspank promoter (IPTG

inducible). The active site of SppA is composed of a catalytic dyad, between a serine (S147) of one

SppA protomer and a lysine (K199) of the adjacent protomer. To obtain an inactive form of SppA, we

mutated the codon for the Lys199 changing it to a codon for alanine (K199A), as described in (Nam,

Kim, and Paetzel 2012). The final strain was named GHBs 088 (BSB1 amyE::PhssppAK199A /spec), and

to simplify Phs sppAK199A.

Figure 52 shows the cell morphology expected for BSB1 without (Figure 52A) and with

(Figure 52B) IPTG: normal B.subtilis rod-shaped cells, while Phs sppA showed damaged long cells.

Interestingly, Phs sppAK199A (Figure 52B) cells did not show a cell growth defect but the normal cell

morphology observed for the BSB1 strain. These results allowed us to conclude that the cell

morphology defect is not a consequence of overexpressing a membrane protein but that the

morphology defect was due to the protease activity of SppA.

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Figure 51. Cell morphology of the sppA, yteJ or sppA yteJ overexpression mutants. The strains, BSB1, Phs

sppA, Phs yteJ and Phs sppA yteJ were grown overnight in LB at 37°C, supplemented with 5 μg/ml Cm and 100

µg/ml Spec when required, and diluted 400-fold into fresh LB medium. Then samples were collected during the

early stationary (stat) phase for microscopy analysis. Phase contrast microscopy and fluorescence microscopy

pictures are shown. Membranes (false coloured red) were stained with FM4-64 and DNA (false coloured blue)

were stained with DAPI. Scale bar is 10 micron.

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103

Figure 52. Cell morphology of the sppA and sppAK199A overexpression mutants. The strains, control strain

(BSB1), PhssppA and PhssppAK199A were grown overnight in LB at 37°C, supplemented with 100 µg/ml Spec

when required. The next day the strains were diluted 400-fold into (A) LB and (B) LB supplemented with

100 µM IPTG. Then samples were collected during the early stationary (stat) phase for microscopy analysis

(Phase contrast microscopy pictures). Scale bar is 10 micron.

2.2.9 SppA and YteJ cell localization.

To better characterize the SppA/YteJ complex, we tried to study the cell localization of these

two proteins. We constructed two strains expressing a translation fusion of each gene, sppA and yteJ,

with sfGFP, by insertion at the amyE locus under the control of the hyperspank promoter. These

constructs are described in the material and methods section 4.3.4. These constructs were confirmed

by PCR and sequencing. The final strains are GHBs 064 (BSB1 amyE::PhsyteJ-sfgfp /spec), GHBs 069

(BSB1 ∆sppA::cat amyE::Phs sfgfp-sppA /spec). For the sake of clarity, strains GHBs 064 and 069 will

be named YteJ-sfGFP and sfGFP-SppA, respectively.

In Figure 53, for both strains YteJ-sfGFP and sfGFP-SppA, the sfGFP signal was found

mostly at the cell septum. However, both proteins showed a cytoplasmic signal too, perhaps due to

problems of translocation of the fused proteins. sfGFP-SppA was observed in foci around the cell

membrane or poles. These results indicated that this protein complex was possibly located at the cell

septum, but our observations did not allow us to conclude with a high level of confidence.

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Figure 53. SppA and YteJ cell localization. The strains, YteJ-sfGFP and sfGFP-SppA, contain a sfGFP

translational fusion with yteJ and sppA genes, respectively. Strains were grown in LB at 37°C; samples were

collected during the early stationary (stat) phase for microscopy analysis. Phase contrast microscopy and

fluorescence microscopy pictures are shown. sfGFP is false coloured of green. Scale bar is 10 micron.

2.2.10 SppA and YteJ in vitro purification.

To analyse the role of YteJ in the complex and the SppA protease activity in vitro, we

produced and purified B. subtilis SppA, YteJ and SppA/YteJ (from an artificial operon) from E. coli.

For that, from the pJ411 plasmid, we constructed plasmids harbouring a copy of each gene or the full

operon fused to a His-tag sequence, under the control of the IPTG-inducible pT7 promoter, and a

kanamycin resistance cassette. These plasmids were used to transform E. coli. The final genes

sequences on these plasmids were confirmed by PCR and sequencing. The his-tagged proteins were

purified as described on the section 4.9.6 of material and method. To simplify, we will refer to the his-

tagged purified proteins as SppA, YteJ and SppA/YteJ.

Purifying active membrane proteins is a challenging task, as it means extracting proteins from

the membrane maintaining its native environment to allow the proteins to preserve its fully folded

conformation. This process can destabilize the protein structure impairing its activity. A strategy to

avoid this problem and study a membrane protein activity is to purify the protein without its

transmembrane domains, transforming the membrane protein in a soluble protein that can be recovered

from the cell cytoplasm. This strategy was used by Nam et al., 2012, which allowed to obtain the

crystallographic structure of SppA and discover that this protein is able to digest fully folded proteins

(Nam, Kim, and Paetzel 2012; Nam and Paetzel 2013).

Chapter 2.RESULTS

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Figure 54. Scheme representing the different His-tag constructions. The rectangles represents SppA and YteJ

proteins, orange represents cytosolic domain, green the transmembrane domains and blue extracellular domain.

The numbers represent the length in amino-acids.

As such, we constructed and purified, as described by Nam et al., 2012 , a SppA truncated of

its transmembrane domain, the first 52 amino acids from SppA N-terminus. This protein was

overexpressed and purified from E. coli cytoplasm and the protein was named SppA∆1-52. In this study,

we used SppA∆1-52 as a control for SppA activity. The purified SppA∆1-52 (lane 2, Figure 55A) showed

an expected band at around 28 kDa. In the case of the full length SppA (lane 3), we observed at least

three bands, one at 36 kDa which was expected as the theoretical molecular weigth (MW) is 36.6 kDa

and two lower bands. These bands of lower MW may indicate two different forms of SppA might

result from a partial autoprocessing of the C-terminal domain of SppA. The purified YteJ (lane 2,

Figure 55B) shows a band of the expected size for YteJ-His, around 20 kDa. When SppA is expressed

together with YteJ-His (lane 4, Figure 55A), SppA (highest band ≈36 kDa) was pulled down with

YteJ-His (lowest band ≈20 kDa), indicating that these two proteins formed a membrane protein

complex even when produced in E.coli.. Comparing with the full length SppA sample (lane 3), the

SppA in the SppA/YteJ complex (lane 4) – i.e. pulled down with the YteJ-His, was mainly observed as

a single band. This single band corresponded to the highest band of SppA full length (≈36 kDa). In

addition, all the full length proteins, purified from the E. coli membrane solubilized with DDM

showed additional faint bands that corresponded to contaminants.

SppA has been described to form an octomer (Nam, Kim, and Paetzel 2012), a structure

obtained by crystallography of SppA∆1-52. We wondered whether this octomeric structure could also be

that of the full length membrane SppA. The same question applied toYteJ. To get rid of contaminants

we performed a second setp of purification: a gel filtration using a Superdex 200 HR 10/300 GL

columnthat allows the separation of proteins from 10 up to 600 kDa. The curves and SDS-PAGE gels

showing the gel filtration results can be found Figure S 2. These results showed that SppA possibly

forms an octomer around 300 kDa, as described in the literature, and YteJ may form an octomer

around 160 kDa.

When loaded on the gel filtration column, the complex SppA/YteJ dissociated as the two

proteins were eluted separately from the column, as two separated octomers. This result indicated that,

Chapter 2.RESULTS

106

in the conditions used for the gel filtration experiment, the individual complex of each protein, SppA

and YteJ, can maintain their structure complex. However, the interaction between the two YteJ and

SppA octomers appeared to be weaker than the intra-protein interactions. One possible explanation for

this weakness is that the complex SppA/YteJ could be made of two subunits, each subunit is an

octomer of each protein, and that the dissociation between the subunits can occur.

Figure 55. SDS PAGE analysis of SppA∆1-52, SppA, SppA/YteJ (A) and YteJ His-tagged (B) purified

proteins. Analysis of the purified samples on 12.5% SDS-PAGE gels, coomassie blue stained. For both gels, A

and B, the first lane corresponds to protein markers, which the bands correspondent to 40, 30, 25 and 20 kDa are

highlighted.

2.2.11 SppA can digest fully folded proteins or not

In 2012, Nam and its colleagues showed that the active soluble SppA was able to digest fully

folded proteins, based on one example, the E.coli lipo-protein BamD (Nam, Kim, and Paetzel 2012).

To test the SppA∆1-52 activity we mixed it with different proteins, available in our laboratory, and

incubated overnight at 37°C in the same buffer conditions as described by Nam et al., 2012.

Interestingly, SppA∆1-52 was not able to digest the bovine serum albumin protein (BSA) and lysozyme

(Figure 56A lanes 2, 3, 6 and 7, respectively), as the band intensities remained the same for the BSA

or lysozyme with or without the presence of SppA∆1-52. However, in the case of GFP we can see a shift

on the size of the core band when treated with SppA∆1-52. This GFP, hereafter called GFP-ext, was

modified by the addition of a around 3 kDa C-terminal tail (24 amino acids, usually used as a linker

Chapter 2.RESULTS

107

for GFP fusion). The shift on the band size corresponded to the digestion of this 3 kDa tail, the core of

the GFP remaining intact. We then tested different proteins, four are shown on Figure 56B. SppA∆1-52

could not digest the E. coli DNA single strand binding protein (SSB) (lane 3). When treated with

SppA∆1-52, the bands corresponding to the following proteins disappeared: B. subtilis nucleoid

associated protein (Rok) and E. coli DNA-binding protein (Ec-HNS). These two proteins were

completely digested (Figure 56B, lane 5 and 7), therefore we aslo proved that SppA can digest fully

folded proteins. However, the B. subtilis catabolite control protein A (CcpA), when mixed overnight

with SppA, disappeared as a whole protein leaving a lower MW band still present. This is an example

of a partial digestion of a protein by SppA∆1-52 (lane 9). These results allowed us to conclude that, as

described in the literature, SppA∆1-52 can digest fully folded proteins. However, as the digestion was

sometimes not possible for some substrates, occured sometimes only partially for others, it is highly

probable that the protease activity of SppA∆1-52.depends on the structure of the substrate.

Figure 56. SDS PAGE analysis of SppA∆1-52 activity. For both gels, (A) and (B), the first lane corresponds to

protein markers, for which the bands corresponding to 40 and 25 kDa are indicated. SppA∆1-52 alone at the last

right lane of gel A.. Protein substrates: bovine serum albumin (BSA), GFPext, lysozyme, E. coli DNA single

strand binding protein (SSB), B. subtilis nucleoid associated protein (Rok), E. coli DNA-binding protein (Ec-

HNS) and B. subtilis catabolite control protein A (CcpA). (-) protein substrate alone and (+) SppA∆1-52 and the

substrate protein. The samples were incubated 16 h at 37°C.

2.2.12 YteJ inhibits SppA activity

Based on the previous results we decided to use the GFP containing C-terminal unstructured

tail as substrate to study the protease activity of SppA as well as SppA in the SppA/YteJ complex.

Their activity were tested overnight at 37°C (Figure 57A). SppA∆1-52 and SppA were able to digest the

substrate, as a MW downshift of the GFPext was observed. However, when in complex with YteJ and

mixed with the GFPext substrate, SppA did not completely digest the substrate, as two bands

corresponding to both, GFP and GFPext, were visible on the gel. Altogether these results revealed the

negative effect of the presence of YteJ on the activity of SppA.

Chapter 2.RESULTS

108

As it seemed that the activity of SppA was slower when SppA was in complex with YteJ, we

performed more precise kinetic experiments following the protease activity of the different forms of

SppA by taking samples at intervals of one hour at 37°C. As the Bradford assay to determine the

concentration of SppA was not very accurate with membrane proteins, we chose to estimate the

quantities of SppA for our “different” SppAs on SDS-PAGE. The band intensities were used to adjust

the quantity of SppAs mixed with the GFPext substrate.

Figure 57. SDS PAGE analysis of SppA∆1-52, SppA and SppA/YteJ activity. A and B corresponds to 12.5%

SDS-PAGE gels, coomassie blue stained. A. activity after 16h at 37°C; B. kinetics of 60 minutes of activity at

37°C. C. Graph of protease activity kinetics of the different forms of SppA. The intensity of the product bands

were measured by Image Lab (Biorad software).

In Figure 57B and C, after 15 min SppA∆1-52 was able to digest almost 100% of the substrate,

while SppA was only able to digest 50% of the substrate after 60 min. The protease activity of the full

length membrane SppA appeared therefore lower than that of the soluble SppA. The presence of the

transmembrane domain of SppA might be involved in the regulation of SppA activity. For one hour, in

complex with YteJ, SppA was not able to digest the substrate. This result indicated that YteJ may act

as a negative regulator of SppA activity.

An active SppA is only purified without its C-terminal, that the enzyme self-processes as

described in (Nam and Paetzel 2013). The authors also showed that an inactive SppA, SppAK199A, was

purified with its unprocessed C-terminal domain.

Chapter 2.RESULTS

109

On our SDS-PAGE analysis, SppA appeared as three bands (lane 2, Figure 58A), while the

inactive SppAK199A appeared as a single band, whose MW was higher than that of the active SppA

(lane 3). When SppA is expressed together with YteJ, we observed a band for SppA which was higher

than active form of SppA but slightly lower than the SppAK199A inactive form. The hypothesis was

that:

1) when active SppA is expressed and purified alone, it can autocatalitycally remove its own

C-terminal domain and appeared as multiple bands on gel (different stages of self-processing of the C-

terminus)

2) SppA in the SppA/YteJ complex probably still contains its C-terminus and appears as a

higher MW band comparable to that of the inactive SppAK119A.

To confirm this hypothesis we performed a MS/MS analysis of the major bands for SppA and

SppA+YteJ shown on Figure 58A. The results showed that (i) the 3 bands of lane 2 were SppA,

although the peptides from the C-terminal could not be detected (ii) for SppA purified in complex with

YteJ, two peptides belonging to the C-terminal domain of SppA were present (peptides are highlighted

in Figure 58B (highlighted by green and yellow). (iii) finally the second major band on lane 4 was

YteJ. We therefore concluded that the presence of YteJ inhibits the SppA-mediated self-cleavage of its

C-terminal part. The presence of a normally absent C-terminal domain in an active SppA indicated

that SppA was inactive in the complex SppA/YteJ, and it confirmed the results obtained with the

enzymatic experiments shown in Figure 57.

Figure 58. SDS-PAGE and mass spectrometry analysis of SppA and SppA/YteJ complex (A) The first lane

corresponds to protein marker, for which the band corresponding to 40 kDa is highlighted. All the proteins

contain a his-tag. Expected bands for SppAK199A and SppA≈37 kDa and YteJ 19 kDa. (B) SppA protein

sequence. The major bands of SppA and SppA/YteJ were analysed by MS/MS as indicated in the figure with

labels i, ii, iii. The MS/MS analysis showed that the band of SppA, in the SppA/YteJ sample, contains two

peptides belonging to the C-terminal domain of SppA (highlighted in green and yellow).

Chapter 2.RESULTS

110

2.2.13 The C-terminal domain of YteJ is involved in the regulation of SppA

activity

Following the previous results, we wanted to understand how YteJ is able to inhibit the SppA

activity in vivo. To try to understand which domains of YteJ blocks SppA activity we used the

morphological defective phenotype, caused by the overexpression of SppA, as a tool. With that

objective we have constructed new mutant strains overexpressing sppA yteJ, with different truncated

versions of YteJ and analyzed their cell morphology and compared with Phs sppA strain (Figure 59 A).

Therefore, when overexpressing these constructs if YteJ is still able to inhibit SppA we should expect

normal cell like BSB1 wild type strains, by other hand if we truncate the domain that blocks SppA we

should expect to see long filamentous cells with division defects. Based in its secondary protein

structure, the C-terminal of YteJ was deleted with three different lengths. In the first strain, YteJ was

truncated from its short C-terminal α-helix corresponding to the last twelve amino acids (Figure 59

B). Secondly, we have truncated YteJ from thirty five amino-acids from its C-terminal region (Figure

59C). At the last construct we have deleted the full C-terminal domain and substituted by the SPA tag

(Figure 59 D). These constructs were re-inserted at the amyE ectopic locus, into the BSB1 wild type

background, under the control of the hyperspank promoter, an IPTG inducible promoter. These

constructions included their natural RBS and a spectinomycin (Spec) resistance cassette. These

constructs were confirmed by PCR and sequencing. The final strains are GHBs 085 (BSB1

amyE::Phyper-spank sppA yteJ ∆129-164 spec), GHBs 087 (BSB1 amyE::Phyper-spank sppA yteJ ∆152-164

spec), GHBs 088 (BSB1 amyE::Phyper-spank sppA yteJ ∆92-164SPA spec). In order to simplify the

understanding, from here, the strains GHBs 085, 087 and 088 will be named Phs(sppA-yteJ∆129-164),

Phs(sppA-yteJ∆152-164) and Phs(sppA-yteJ∆92-164SPA), respectively.

Figure 59. Scheme of SppA/YteJ protein complex in the different YteJ truncated constructions. (A) sppA

yteJ overexpression strain; (B) sppA yteJ ∆152-164 overexpression strain, the las 12 amino-acids from YteJ C-

terminal were removed; (C) sppA yteJ ∆129-164 overexpression strain, truncated YteJ from thirty five amino-acids

from its C-terminal region; (D) sppA yteJ ∆92-164 overexpression strain, the C-terminal domain was removed and

substituted by a SPA-tag; Orange represents cytosolic domain, green the transmembrane domains and blue

extracellular domain. The numbers represent the length in amino acids.

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111

Figure 60. Cell morphology of the overexpressing strains. All the strains grown in LB until early stationary

phase, then a sample was taken for microscopy analysis. The first panel (left side) shows the phase contrast

microscopy pictures of un-induced strains, indicated genotypes. The remaining panels (from left to right side)

represent the phase contrast and fluorescence microscopy of the indicated strains grown in IPTG presence. sppA,

sppA yteJ, sppA yteJ∆152-164, sppA yteJ∆129-164 and sppA yteJ∆92-164SPA were expressed from an artificial Phs

promoter and induced with 50µM IPTG. Membranes (false coloured red) were stained with FM4-64 and

nucleoids (false coloured blue) were stained with DAPI. Scale bar is 10 micron. Black arrows represents the cell

size, by indicating the end and begging of the cell; White triangles represents the cell septum or septum

formation points.

Overnight cultures were diluted 400-fold into fresh pre-warmed LB media without any

supplements. Biomass (OD600) was measured at regular intervals and samples were collected during

the early stationary (stat) phase of the growth cycle and observed by microscopy, as described at the

previous section. Cells were stained with the membrane dye FM4-64, allowing the observation of the

cell membranes and septum, and with DAPI, allowing the observation of chromosomal segregation.

The cells of the BSB1 strain (Figure 60-first line on the panel) appeared to be normal B. subtilis rod-

shaped cells, well-defined division septa, cell poles and well segregated chromosomes. The

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112

overexpression of sppA affected the cell morphology creating elongated cells, as the observed in our

previous results (second line Figure 60). Surprisingly, when we overexpressed sppA-yteJ∆152-164

(fourth line Figure 60), YteJ with the short truncation at its C-terminal, we could observe an effect in

the cell morphology. Even though the cells were not so long as PhssppA cells, the cell were longer

than the BSB1 cells, with an evident septa mal formation and an abnormal chromosomal segregation.

Furthermore, the overexpressed sppA-yteJ∆129-164 (fifth line Figure 60) restored the long phenotype

observed for the sppA overexpression. Contrarily, the strain containing a swap between the C-terminal

region of YteJ by the SPA-tag (sixth line Figure 60), Phs (sppA-yteJ∆92-164SPA), show shorter cells

similar to the BSB1 and PhssppA yteJ cells (first and third lines Figure 60). However, these cells

show cell wall and division problems. These results allowed us to conclude that the C-terminal domain

of YteJ is involved and important for the regulation of SppA activity, as the removal of the last 25

amino acids affected the cell growth, and that removing the last 45 amino acids of YteJ led to a a cell

phenotype similar to what was observed with the overexpression of SppA alone, i.e. a complete

deregulation.of SppA protease activity.

2.2.14 SppA digests subtilin

Our previous results showed that SppA∆1-52 can digest fully folded proteins, depending on the structure

of the substrate. In this chapter of results we have also demonstrated that SppA have an important role

in the lantibiotic resistance mechanism. Assuming that the lantibiotics are peptides, we were

wondering if in this resistance mechanism SppA is directly digesting the lantibiotics. As such we used

the previous purified and active SppA∆1-52 to test our hypothesis. SppA∆1-52 was incubated overnight in

a subtilin-containing reaction buffer (20 mM Tris-HCl pH7.5, 200mM NaCl) at 37°C, as well as the

controls, SppA∆1-52 alone in the reaction buffer, subtilin solution and a positive control with proteinase

K. Proteinase K is a broad range serine protease, the same family that SppA (MORIHARA and

TSUZUK 1975). Then these compounds were tested with the wild type BSB1 and ΔsppA strain. The

experiment was done as described at section. At Figure 61, the first graph top left of the panel

corresponds to the controls, i.e. the BSB1 and ∆sppA mutant strain grown in LB without subtilin. The

two strains showed identical growth rates. When treated with 30% subtilin (graph top right), the

growth of the strains was affected, with a visible difference in the resistance to subtilin between the

BSB1 and the ∆sppA. ∆sppA took longer to recover its growth and reach an OD600 of 0.1. The subtilin

was stable and did not degrade overnight at 37°C, as it induced the same type of growth delay (graph

middle left), than the fresh subtilin used. When the subtilin was treated overnight with the SppA∆1-52,

we can observe that the growth is similar to the control without subtilin (graph top left), without

differences between the strains. The same was obtained for with the subtilin treated with proteinase K.

With these results we can conclude that the subtilin was degraded by both SppA∆1-52 and proteinase K.

Therefore, we can hypothesize that, in vivo, the direct degradation of subtilin by SppA is part of the

mechanism of action against lantibiotics activity.

Chapter 2.RESULTS

113

Figure 61. Growth of BSB1 and ∆sppA in the presence of different subtilin solutions. The BSB1 and ∆sppA

strains were grown in LB in a 96-well plate, and the OD600 was measured at specific time points. The different

graphs represent the growth of the strains in different conditions: top left in LB without subtilin; top right LB

supplemented with 30% subtilin; in the remaining graphs, the strains grown in 30% subtilin treated 16h at 37 °C

just in reaction buffer (20 mM Tris-HCl pH7.5, 200mM NaCl) or mixed with SppA∆1-52 or proteinase K,

respectively; The conditions are labelled on the top of each graph.

Chapter 2.RESULTS

114

115

DISCUSSION AND

PERSPECTIVES

Chapter 3. DISCUSSION AND PERSPECTIVES

117

3 DISCUSSION AND PERSPECTIVES

The ability to transport newly synthesized proteins across and into the bacterial cytoplasmic

membrane is an essential mechanism to maintain cell integrity. This transportation mechanism

involves universally conserved protein-conducting machineries that can transport a wide variety of

potential substrates. For example, the Sec translocon mediates the translocation of secretory and

membrane proteins in an unfolded conformation. The SecYEG, the core channel of the bacterial Sec

translocon, is the minimal membrane-embedded unit required for protein transport (Collinson 2005).

However, the full translocation process from the recognition of the pre-protein, its translocation and

final maturation involve a dynamic between the protein-channel and associated proteins, which

interact and work together (de Keyzer et al. 2003). Although the mechanisms involved in the Sec

pathway has been extensively studied, the knowledge about B. subtilis protein complexes is limited.

The possible protein-protein interactions within the B. subtilis Sec pathway were studied and will be

discussed in section 3.1. As SppA was initially identified as an accessory protein involved in the Sec

pathway, the section 3.2 will focus on its interaction with YteJ, its role and activity in the cell.

3.1 Protein interactions of the B. subtilis Sec pathway

In E. coli, it has been established that the Sec holo-translocon is a supramolecular complex

composed of the membrane protein complexes, SecYEG (Sec translocon), SecD/SecF, YajC and YidC

(Schulze et al. 2014; Komar et al. 2016; Botte et al. 2016). Based on the multiple interactions found in

E. coli, we intended to study these interactions in B. subtilis.

Our experiments of sequential peptide affinity purifications with the SPA-tagged proteins

SecDF and SpoIIIJ in B. subtilis showed that the B. subtilis SecDF, homologous to a fusion of the E.

coli SecD and SecF, and B. subtilis SpoIIIJ, homologous to E. coli YidC, did not co-purify. According

to these results, it seems that SecDF and SpoIIIJ do not form a stable complex in B. subtilis, unlike

what was shown in E. coli. However, the absence of co-purification of B. subtilis SecDF and SpoIIIJ

does not completely rule out the possibility that these two proteins interact. Their interaction can be

weaker than that in E. coli, and might have been abolished by the use of a too high concentration of

detergent, the n-dodecyl β-D-maltoside (DDM). In our procedure, we classically used a DDM

concentration of 1% which might affect protein interactions, although 2% DDM concentration was

used in E. coli to solubilize the membranes. We therefore tested a range of different DDM

concentrations, from 0.5% to 2%, and also reduced the purification steps to only one affinity column

to avoid losing some protein partners. Analysed by LC-MS/MS, the results of these experiments

showed that the SpoIIIJ did not co-purified with the SecDF-SPA protein. However, in the reciprocal

experiment, after affinity purification of SpoIIIJ-SPA, low amounts of SecDF were detected regardless


118

of the DDM concentrations. This suggested that B. subtilis SecDF and SpoIIIJ may eventually interact.

If so, this interaction might be rather weak as the addition of a second affinity column had previously

abolished it. Moreover, as this result was only obtained when the SPA tag was added to SpoIIIJ, it also

suggests that the SPA tag fused to SecDF might have precluded the interaction. However, in

conclusion, if B. subtilis SecDF and SpoIIIJ interact, these two proteins do not form a stable complex

like in E. coli.

Unfortunately, in our attempt to study the translocon SecYEG, B. subtilis strains expressing

SecY-SPA and SecE-SPA could not be obtained. Therefore we analysed only B. subtilis SecG-SPA by

tandem affinity purification. It turned out that SecG-SPA did not co-purify with the expected SecY or

SecE, nor SecDF, unlike what had been observed in E. coli. Again, we cannot rule out the possibility

that the SPA tag fused to SecG have induced a dissociation of the other proteins, SecY and SecE, or

weakened their interactions. However, it is highly unlikely as the translocon, SecYEG, is essential to

the cell and as any defect in the formation of this complex should harm the cell. Our result rather

reflects the fact that the translocon, SecYEG, or the holotranslocon, SecYEG-SecDF-SpoIIIJ, might

not be a very stable protein complex in B. subtilis.

Combined together, these results suggest that B. subtilis do not possess a stable holo-

translocon as the E. coli SecYEG-SecDF-YajC-YidC supracomplex. In E. coli the existence of this

holo-translocon as a supracomplex was shown by different methods including BN-PAGE and the

direct interactions within the proteins forming the holotranslocon were further refined by crosslinking

experiments (Schulze et al. 2014; Komar et al. 2016). To study the “holotranslocon” in B. subtilis,

cross-linkers should be used and added to the cells before purification of any of the translocon

components.

After translocation, the secretory proteins are released from the membrane and undergo a

maturation process carried out by other accessory proteins to the Sec machinery, as for example the

thiol-disulfide oxido-reductases (Bdb proteins), which during protein folding, are responsible for

rearranging incorrect disulfide bonds (Tjalsma et al. 2004; Driessen & Nouwen 2008). Structural

studies showed that the E. coli thiol-disulfide oxido-reductases DsbC and DsbD form a complex.

DsbD binds to the central cleft of the V-shaped DsbC dimer causing a shift into a close conformation

and it activates DsbC (Fabianek et al. 2000; Haebel et al. 2002). We also investigated this connection

between the B. subtilis BdbB and BdbD, two thiol-disulfide oxido-reductases homologous to E. coli

DsbC and DsbD. We did not find evidences that these two proteins form a complex either, as our

results showed that, when SPA-tagged, BdbB or BdbD did not co-purify with each other.

3.2 SppA/YteJ membrane protein complex

As shown in Chapter II of the results, we have found evidences of a stable complex between

the B. subtilis SppA, the signal peptide peptidase, and the unknown membrane protein YteJ.

119

The first evidence for the existence of this complex was obtained by BN-PAGE experiments,

which showed that these two proteins co-migrated together, suggesting that they might interact. The

tandem affinity purification of YteJ-SPA showed that SppA was co-purified with YteJ-SPA,

indicating again that these two proteins interact and form a stable complex. For the in vitro study,

when the his6-sppA yteJ operon was expressed in E. coli, the purification of the his-tagged SppA

clearly showed the presence of YteJ, reinforcing the idea that a membrane protein complex made of B.

subtilis SppA and YteJ can be formed in E. coli.

The existence of this complex in the cytoplasmic membrane of B. subtilis prompted us to

study whether YteJ could play a role in this complex. I will discuss our results in the following order:

1. the role of SppA in protein secretion ?

2. the role of SppA (and YteJ) in the resistance to nisin, subtilin and LP9

3. the protease activity of SppA is regulated by YteJ

4. SppA/YteJ, a complex involved in cell division ?

5. YteJ, another function?

3.2.1 Role of SppA and YteJ in protein secretion:

In 1999, Bolhuis et al. observed that the deletion of sppA induced a slight decrease in the

secretion of mature secretory proteins, and more importantly a delay in the maturation process of the

hypersecreted protein AmyQ. As SppA was believed to be involved in the degradation of the signal

peptides of the secreted proteins, this result was interpreted as a consequence of the presence of

remnant signal peptides in the membrane, that would cause a slow-down of the processing of the pre-

proteins under hyper-secreting conditions. Here, in this study, we have shown that the deletion of sppA

and yteJ do not affect the secretion of a highly secreted protein, AmyM. However, Bolhuis et al.

(1999) used pulse chase experiments to show the maturation delay of the hypersecreted AmyQ.

Therefore, our deletion strains were also tested by pulse chase experiments (see Figure S 3), in Prof.

van Dijl’s laboratory (Groningen, The Neterlands). Their results showed no major effect of any

deletion (ΔsppA, ΔyteJ or Δsppa yteJ) on the secretion of the model protein AmyM. Taking into

account these results, our conclusions are that neither SppA nor YteJ are directly involved in protein

secretion. It does not rule out any indirect effect that could explain what was previously observed.

The results obtained with the SppA, or YteJ overexpression strains, overexpressing, showed a

negative effect on the AmyM secretion level when SppA is overexpressed, while the overexpression

of YteJ, or SppA and YteJ together, did not lead to any difference. This indicated that, SppA, able to

degrade secreted proteins, could be detrimental when overexpressed for the production of secreted

proteins. Our result is in complete disagreement with previous results in Bacillus licheniformis for

which the overexpression of SppA proved to be beneficial for the secretion levels of nattokinase and


120

AmyL (Cai et al. 2017). In this study, sppA was under the control of the constitutive promoter P43, a

promoter which is thought to be active during the lag and exponential phase of growth but not during

the stationary phase. In our study, we used the promoter Phs induced by the addition of IPTG, therefore

SppA was expressed, regardless of the growth phase. This might explain the difference of results

between the two studies, as in both cases, the extracellular secreted protein contents were assessed in

late stationary phase. In conclusion, the role of SppA in protein secretion appeared to be quite limited

as worst negative effects of 20 % or best positive effects of 60% in the protein secretion levels were

reported (Bolhuis et al. 1999)(Cai et al. 2017), as opposed to the positive effects of 250-1000 %

obtained after the overexpression of RasP (Neef et al. 2017).

3.2.2 Role in the resistance to nisin, subtilin and LP9

The role of SppA was more obvious in the resistance to nisin, subtilin and LP9 as the deletion

of sppA induced a higher sensitivity of the cells to these antibiotic peptides. These results were in

agreement with previous observations that showed that sppA, a member of the σW regulon, was

involved in the lantibiotic resistance (Kingston, Liao, and Helmann 2013b).

Our experiments with the sppA complementation strain confirmed the direct role of SppA in

this resistance as the complemented cells showed a lantiobic or LP9 resistance identical to the wild-

type cells. We also tested the direct digestion of subtilin by SppA by incubating the purified SppA in a

subtilin-containing LB medium for 12 hours at 37°C, and we observed that the ΔsppA cells grew as

well as the wild-type cells in this SppA-treated medium (Figure 61).

The role of YteJ appeared more complicated to understand with the antibiotic peptide

experiments. The observed phenotypes of the ΔyteJ cells showed either a slightly higher sensitivity to

the antibiotic peptides than the wild-type cells, or a greater resistance. It seemed to be dependent on

the concentrations of the peptide used. If the resistance to the lantibiotic peptides or LP9 is only due to

the protease activity of SppA, the ΔyteJ phenotype might be explained by the SppA regulatory role of

YteJ. We showed that YteJ negatively regulates the activity of SppA; however, YteJ has to release its

inhibitory action on SppA, presumably by responding to a signal, induced by the antibiotic peptides. In

the absence of YteJ, the activity of SppA might not be timely regulated, hence the lower antibiotic

resistance. This hypothesis could not account for the higher antibiotic resistance observed in ΔyteJ

cells in some cases.

3.2.3 Regulation of the sppA yteJ operon

In parallel with the antibiotic resistance study, we also studied the transcription regulation of

the operon sppA yteJ. In Nicolas et al (2012), a second promoter, σA-dependent, was detected

upstream of the σW-dependent promoter. Our experiments using sGFP fusions with one or potentially

two promoters showed that the major promoter was the σW-dependent promoter. As such, we

confirmed that sppA and yteJ are part of the σW regulon, a regulon that is expressed when cells are

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subjected to an envelope or cell wall stress. Interestingly, the operon structure of these two genes,

sppA and yteJ, is well conserved across the bacilli class, which includes Bacillus subtilis but also

Listeria monocytogenes.

3.2.4 SppA/YteJ, a complex involved in quality control of cell division?

As shown in the chapter II of the results, we observed that high levels of expression of SppA

create a severe cell growth defect, giving rise to a filamentous phenotype (see Figure 51). This

phenotype could be due to the overexpression of a membrane protein. The overexpression of

membrane proteins and proteases normally result in the accumulation of cytoplasmic aggregates

containing the overexpressed proteins, accumulation of chaperones (e.g. DnaK), soluble proteases (e.g.

ClpXP) or precursors of secreted and membrane proteins (Wagner et al., 2007). To discard this

hypothesis, we constructed a strain that overexpressed an inactive form of SppA, the SppAK199A. The

analysis of the morphology of the cells showed a normal cell morphology (see Figure 52). These

results allowed us to conclude that the activity of SppA is responsible for the cell growth defect. In

Figure 51 the affected long cells of the PhssppA strain are not just only filamentous but they seem to

show a defect on cell division, malformation of the septum and impaired chromosomal segregation.

This morphological phenotype defect is reminiscent of the cell defect phenotype described in deletion

mutants involving protein regulators of the divisome. The B. subtilis cell division is well coordinated

with the elongation, as it requires a switch in the orientation of the cell wall synthesis, and the

formation of the mid-cell Z ring for cell constriction and division (Adams and Errington, 2009; Leaver

et al., 2009; Errington and Wu, 2017). The Min system prevents the polar division of the cells. It was

shown that the deletion of B. subtilis noc combined with the depletion of MinD lead to a defect in the

division, absence of division septa formation and elongated filaments phenotype, similar to what we

observed on our results obtained for the overexpression of SppA (Figure Figure 51 and Figure 60) (L.

J. Wu and Errington 2004). It was shown too that the B. subtilis minJ deletion mutant, comparing with

the wild type strain, grew as long filaments of cells with sporadic septa (Figure 62), as we observed in

our mutants (Patrick and Kearns 2008). However, in both cases the segregation of the chromosomal

DNA was not affected, as it is possible to observe a well divided nucleoids on Figure 62, the contrary

of our results, that showed long filamentous cells with a unique long nucleoid (Figure 51).


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Figure 62. Cells mutated for minJ are filamentous. Fluorescence microscopy of wild type (A), ∆minJ (B)

were grown to mid-exponential phase and stained with FM 4-64 (membrane, false coloured red) and DAPI

(chromosome, false coloured blue). Scale bar is 4 mm. (Patrick and Kearns 2008).

Other types of proteins could be the targets of an overexpressed, active and unregulated SppA,

as for example the penicillin binding proteins (Pbp). Mutants of some Pbps show similar phenotype of

elongated cells as well (Figure 63).

Figure 63. Morphology of B. subtilis cells lacking multiple penicillin-binding proteins (Pbp). (A) wild-type

strain; (B) cells lacking Pbp1; (C) cells lacking Pbp1 and 4; (D) cells lacking (Adapted from Popham and Setlow

1996)).

In conclusion, the clear cell morphology phenotype defect observed in a strain overexpressing

an active SppA points out that proteins involved in the division process or cell wall synthesis are

affected. However, no specific substrate for SppA can be inferred from this observed phenotype, as

too many proteins can be involved.

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3.2.5 The protease activity of SppA and its regulation by YteJ

3.2.5.1 The protease activity itself:

Nam et al. (2012) performed in vitro studies with a soluble form, truncated B. subtilis SppAΔ2-

52, deleted from its transmembrane domain (amino acids from position 2 to 52). These authors showed

that SppA∆2-54 could digest signal peptides as well as fully folded proteins. Their conclusion was based

on their assay of a successful overnight digestion of a fully folded E. coli lipoprotein (BamD) (Nam,

Kim, and Paetzel 2012). In our study, we have shown that B. subtilis SppA∆2-54 is not able to digest all

fully folded proteins, as we tested SppA activity on seven different proteins available in the lab as

shown in Figure 53. Our study of the activity of SppA∆2-54 with more proteins indicates a new feature

about the enzymatic properties of SppA. As some proteins showed a SppA proteolytically-resistant

domain, our hypothesis is that some protein structures or specific sequences of amino-acids might

have the ability to resist to the proteolytic activity of SppA. To be demonstrated this hypothesis would

require a specific experimental setting involving, for example, a synthetic peptide that includes a

variety of structures or amino acid sequences.

3.2.5.2 The complex SppA/YteJ and its activity:

In vitro, a soluble SppA, truncated from its N-terminal transmembrane domain, SppA∆2-54, has

been crystallized as an octomer (Nam, Kim, and Paetzel 2012). In our hands, the full-length SppA,

expressed and purified from the E. coli membrane fraction, is also an octomer. Its partner, B. subtilis

YteJ contains two to three transmembrane spanning domains, depending on the prediction structural

programs. YteJ was purified as an apparent molecular weight corresponding to an octomer as well.

The results from the two SPA purification experiments showed that the ratio was more probably a 1:1

between SppA and YteJ, which would be in accordance with the fact that the two genes are in operon.

It would imply that the SppA/YteJ complex would be made of two octomers of each protein. Although

the two proteins were purified as a complex from the sequential affinity purification, in which each

protein was fused to a different tag, the subsequent gel filtration experiment showed a complete

dissociation of the two proteins during the chromatography, suggesting a relative weakly interaction

between the two octomers. This interaction between multimers of different proteins is reminiscent of

the membrane protein complex formed by the protease FtsH and HflK and HflC (Ito and Akiyama

2005). FtsH is a hexamer with most of its protease domain facing the cytoplasm. HflK and HflC, two

membrane proteins that associate first as heterodimers, interact with E. coli FtsH, to form a huge

complex of 1 MDa, the FtsH holoenzyme. It was shown that HflKC inhibits the protease activity of

FtsH for one of its substrate, SecY (Kihara, Akiyama, and Ito 1996).

We also showed that, in vitro, in the SppA/YteJ complex, the SppA protease activity was

strongly diminished when compared to that of SppA alone. In vivo, the overexpression of SppA

created morphological defect to the cells, whereas the cell morphology was unchanged compared to

the wild-type cells, when YteJ was co-expressed with SppA. Our conclusion is that YteJ is a regulator


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of the protease activity of SppA. In vivo, we further showed that the last 25 amino acids of the C-

terminal domain of YteJ were involved in this inhibition mechanism, and that removing the last 45

amino acids of YteJ led to a completely unregulated SppA. Nam and Paetzel (2013) had proposed that

an active SppA processes its own C-terminus, and therefore that SppA C-terminus would act as a

negative regulator domain, presumably preventing SppA to become active too early during

translocation. Our findings provide another level of regulation for SppA through its interaction with

YteJ. Our MS data showed that the C-terminal domain of SppA was detectable by mass spectrometry

only when SppA was purified as a complex with YteJ. The two peptides belonging to the C-terminal

domain of SppA that were detected are highlighted in the Figure 58 are 309SEIDFLNMR317 and

318EILSQSGSPR327.

Therefore one could conclude that SppA in the SppA/YteJ complex is maintained in an

inactive form by the presence of YteJ, and SppA in the SppA/YteJ complex is unable to self-process

its C-terminus as an active SppA would do. However, as the SppA last seven amino acids, i.e. the

peptide 328MMYLYAK335, have not been found in our MS analysis, we cannot be certain that SppA,

even in the complex SppA/YteJ, has not self-processed at least part of its C-terminus. Noticeably,

when SppA was purified, it invariably showed multiple bands on SDS-PAGE, at least 3 visible bands,

separated by an apparent molecular weight difference of 3 to 4 kDa, the highest molecular weight band

corresponding exactly to the band of SppA in the SppA/YteJ complex (See Figure 58). This suggested

that SppA, purified alone from the E. coli membrane fraction, was able to self-process its C-terminus

partially, and that this process was slow. On the opposite, Nam and Paetzel (2013) noticed that, for the

N-terminal transmembrane domain truncated SppA, SppA∆2-54, the self-processing of the C-terminus

of SppA was probably very fast and complete in E. coli even before the enzyme was purified. It

suggests that, in our work, the N-terminal transmembrane domain of the full length SppA may also

slow down the self-processing of the C-terminus. Accordingly, the protease activity of the full-length

SppA was slower than that of the soluble form, SppA∆1-52. E. coli FtsH has been shown to self-process

the last seven amino acids of its own C-terminal domain in vivo, that this self-processing was growth

phase-dependent, and that the deletion of hflKC slowed down the C-terminus self-processing of FtsH

(Akiyama 1999). However, for E. coli FtsH, both full length and processed proteins are active. In the

case of B. subtilis SppA,YteJ inhibits SppA by preventing its C-terminus self-processing. By

interacting with SppA, YteJ is able to maintain SppA in its inactive form presumably until a

physiological harm caused by antibiotic peptides requires the activation of SppA. In this view, SppA

activity, that could be detrimental to the cell if not regulated – see next paragraph -, would act only

when necessary.

Finally, this raises the question of the nature of the signal that triggers the activation of SppA

and how this happens. For example, the PDZ domain of E. coli DegS acts as an inhibitory domain of

DegS until it binds hydrophobic peptides of the C-terminal of misfolded proteins leading to the release

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of the inhibition (Walsh et al. 2003). This model of activation could be applied to YteJ and SppA, the

C-terminal domain of YteJ acting as the inhibitory/activating domain for SppA.

3.2.6 YteJ, another function?

YteJ, by analogy to the PDZ domain for DegS, could be viewed as a sensor of a signal

induced when the cell is subjected to a cell wall stress due to antibiotic peptides. The overall YteJ

amino-acid sequence analysis shows that the protein belongs to the RDD superfamily, which is

characterized by the presence of three highly conserved residues, arginine-aspartate-aspartate. This

domain is found in three other proteins in the B. subtilis genome (Yczc, YckC, YxaI), and the Pfam

database indicates that this RDD domain is present in many transmembrane proteins, mainly in

bacteria, but also with some occurrences in Archeae and Eukaryota. It could be the main domain of the

protein or associated with other domains of various functions. Until recently the molecular function of

this RDD domain remained completely elusive. However, looking for novel Na+/H+ antiporters, Shao

et al (2018) identified a protein named RDD in the moderate halophile Halobacillus andaensis. These

authors demonstrated that the RDD protein functions as an Na+ (Li+, K+)/H+ antiporter, but also that

four out of six conserved arginine or aspartate residues were indispensable to the antiporter activity

(Shao et al. 2018). Although B. subtilis YteJ did not appear in the sequence alignment showed by Li

Shao et al (2018), it is clearly related to H. andaensis RDD (Figure 64). Moreover the corresponding

rdd gene was identified adjacent to a putative sppA gene. B. subtilis YteJ shares all six conserved

arginine and aspartate residues identified in the homologs of H. andaensis RDD (Shao et al. 2018),

and overall the two protein sequences show 39% identity.

Figure 64. Amino acid sequence alignment between H. andaensis RDD and B. subtilis YteJ. Conserved

amino acids are highlighted in red.

Therefore one can hypothesize that YteJ might act as a cation antiporter in B. subtilis as in H.

andaensis. In case of salt stress and/or leakage of cations induced by lantibiotic peptides, thought to

create holes in the membrane, YteJ would become more active inducing a conformation change in its

own structure that, in turn, would induce the release of the YteJ C-terminus inhibitory effect on SppA.

The complex SppA/YteJ would be a two-protein system dedicated to fight the lantibiotic effects. In

vivo experimental evidences that B. subtilis YteJ acts as a cation antiporter are needed. Within the B.

subtilis genome, YteJ and its closest related proteins (YxaI, YckC and YczC sharing over 30%

identity) share a comparable predicted structure made of three transmembrane domains and these

proteins belong to the RDD family; however, none of their genes seem to be in operon with another

gene. Therefore, SppA and YteJ might be a unique example of a two-protein system involving a


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possible cation transporter regulating the activity of a protease. In vitro detailed enzyme kinetic studies

combined with structural studies involving mutations in SppA as well as in the C-terminus of YteJ

could reveal how this activating mechanism takes place.

3.2.7 Model for the interaction of SppA and YteJ

We propose the following model in which SppA and YteJ formed a membrane protein

complex in which the two proteins are arranged as two homoactomers bound to each other (Figure

65). Based on our experimental data, SppA in the SppA/YteJ complex would be inactive because YteJ

might interfere with the active site of SppA. In this conformation, SppA could not self-process its own

C-terminal domain, a domain that has been shown to block its active site. Upon a signal, whose nature

remains to be determined, YteJ, probably undergoing a conformational change, would release its

inhibitory effect on SppA that in turn would become active. As lantibiotic peptides are able to create

holes in the cytoplasmic membrane and that YteJ amino acid sequence shows homology with a cation

antiporter of a distant relative of B. subtilis, the nature of the signal that could be sensed by YteJ could

be related to ion homeostasis.

Figure 65. Model for the regulation of SppA by YteJ. (A) shows the SppA² inactive state, where YteJ is

structurally blocking SppA active site; A cell stress (represented by the black icon) induces a cell response that

leads to complex conformational change (represented by a black arrow). This structural modification liberated

the SppA active site, passing to its active state (B).

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3.3 Main results

All of these in vitro and in vivo observations suggest that SppA and YteJ form a strong

membrane protein complex. This study established that YteJ plays a key regulatory role in B.

subtilis SppA activity, highlighting a possible new mechanism of a protein regulating a

protease. The main conclusions of this work are listed below.

BN-PAGE experiments showed us that SppA and YteJ co-migrate together in a native

gradient SDS-PAGE gel, the first evidence of the protein-protein interactions of these

two proteins.

SppA was co-purified with YteJ-SPA.

In B.subtilis, the deletion of sppA affects the resistance to antimicrobial peptides,

while deleting yteJ reveals an intermediate phenotype. The overexpression of SppA

restores the resistance to subtilin.

The high levels of expression of SppA create a cell growth defect and the presence of

YteJ counteracts this effect.

SppA activity is responsible for the cell morphology defect, as when we

overexpressed an inactive version of SppA the cells have a normal phenotype.

In vitro experiments showed that SppA is able to digest fully folded proteins,

depending on their structure.

YteJ inhibit SppA activity, possibly through its C-terminal region.


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MATERIAL AND

METHODS

Chapter 4. MATERIAL AND METHODS

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4 MATERIAL AND METHODS

In this chapter, the molecular biology, microbiology, protein purification and microscopy methods

used throughout this study are described.

4.1 Techniques for DNA manipulation

4.1.1 Oligonucleotide

Oligonucleotides or primers were designed using the Geneious software (Gateway® cloning).

All oligonucleotides, upon arrival, were solubilized in sterilised deionized water and stored at -20°C as

100 pmol/µl stock solution. A detailed list of all primers used in this study is given in Table S1

Appendix.

4.1.2 Polymerase chain reaction (PCR)

The amplification of DNA fragments was done in vitro in by Polymerase Chain Reaction

(PCR). Different DNA polymerases were used according to specific requirements: the High-Fidelity

DNA polymerase Q5TMDNA polymerase (NEB) was used for gene amplification; the

InvitrogenTMPlatinumTMSuperFiTM polymerase was used to amplify long fragments for building

genetic constructs requiring a very accurate replication; and for colony-screening purposes the Thermo

ScientificTMDreamTaq polymerase was used. All the enzymes used on this study were handled

according to the supplier recommendations.

4.1.3 Purification of PCR products

PCR products were purified using the QIAquick PCR Purification Kit (QIAGEN), according

to the manufacturer’s instructions.

4.1.4 Electrophoresis of DNA

The DNA was analysed by electrophoresis of DNA. This technique was performed according

to standard methods using 1% agarose gels and Midori Green Advance (NIPPON Genetics) as

fluorescent dye for detecting DNA.

4.1.5 Purification of DNA from agarose gel

For cloning purposes, after DNA electrophoresis, the fragments of interest were purified from

the agarose gel slices using the QIAquick Gel Extraction Kit (QIAGEN) according to the

manufacturer’s instructions.


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4.1.6 Purification of plasmid DNA

Plasmid DNA was extracted and purified from E. coli or B. subtilis using the QIAprep Spin

Miniprep Kit (QIAGEN), according to manufacturer’s instructions. For the isolation of plasmid DNA

from B. subtilis, lysozyme from chicken egg white (Sigma-Aldrich) was included in the resuspension

buffer of the kit, at a final concentration of 2 mg/ml and the cell suspensions incubated at 37 °C for 15

min. All plasmid DNA was eluted with sterile deionized H2O.

4.1.7 Chromosomal DNA extraction

The genomic DNA of B. subtilis was extracted using the GenElute Bacterial Genomic DNA

(SigmaAldrich) kit, according to supplier’s instructions.

4.1.8 Digestion of DNA with restriction enzymes

The DNA strands were digested by restriction endonucleases (NEB). DNA was digested

from 30 min up to 16 hours and inactivated for 20 min, in the conditions recommended by the

manufacturer and using the buffers provided with the enzymes. The digestion products were either

analysed by DNA electrophoresis, purified from the remaining components of the restriction digestion

reaction, as explained in section 1.3, or directly added to the ligation reaction mix.

4.1.9 Ligation of DNA fragments

Digested DNA fragments were ligated by the T4 DNA ligase (INVITROGEN) in the conditions

recommended by the manufacturer. The ligation reaction volume was typically 20µL and contained

the vector and insert DNA in an approximate molar ratio of 1:3. The ligation mixtures were incubated

at room temperature (RT), from 1h up to an overnight, and then used to transform E. coli.

4.1.10 Gibson assembly technology

This method, developed by Dr. Daniel Gibson and his colleagues, allows assembly of multiple

DNA fragments with overlapping regions in a single-tube isothermal-reaction (Gibson et al. 2009).

Typically, the adjacent fragments were designed to contain a homology region of 20 to 40 base pairs at

their extremities. The amplified fragments were added to a assembly master mix (NEBuilder R HiFi

DNA Assembly Master Mix), which possesses three different enzymatic activities - exonuclease,

DNA polymerase, and DNA ligase activities that work in a single buffer. The exonuclease component

creates single-stranded 3´overhangs that facilitate the annealing of fragments that share

complementarity at one end (overlap region). The DNA polymerase incorporates nucleotides to fill in

any gaps within each annealed fragment. The DNA ligase seals the nicks and covalently bounds the

DNA adjacent segments. In more detail, 10µl of assembly master mix were mixed with 0.02–0.5pmols

of DNA fragments, when up to three fragments are being assembled into a vector, and 0.2–1.0pmoles

of DNA fragments when from four to six fragments are being assembled, as recommended by the


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supplier’s. The reaction mixture was incubated at 50◦C for 15 min, when two or three fragments were

being assembled, or 60 min, when more fragments were assembled. The total volume of the resulting

product was then transformed into competent E. coli MC1061 strains.

4.1.11 Ligation-Independent Cloning (LIC)

Ligation-independent cloning (LIC) is a method that allows the ligation of a PCR fragment to

a plasmid without using ligase or an restriction enzyme (Botella et al. 2010). First the The pMUTIN-

SPALIC vector was linearized by digestion with the restriction enzyme AscI and treated with T4 DNA

polymerase. T4 polymerase has a exonuclease activity 3’→5’ that creates the cohesive extremities in

the presence of dATP for the vector, creating a linearized vector with ssDNA extremities.

Oligonucleotides used to amplify the insert (400bp of the end of the gene of interest) were constructed

in order to contain specific extensions, LIC-P1 and LIC-P2 (Fig.57-A) respectively, which are

homologous with the pMUTIN-SPALIC extremities. The insert weas treated with T4 polymerase too,

creating the ssDNA extremities. The two products were then annealed at room temperature with each

other to form a new pMUTIN-SPA plasmid with the PCR fragment inserted upstream of the SPA

sequence. The obtained plasmid is sufficiently stable for direct transformation into competent E. coli

MC1061 strains. The extracted product plasmid was then transformed into B. subtilis and inserted into

the chromosome by single cross-over (Fig. 57-B). pMUTIN-SPA has a Pspac inducible promotor to

assure that after its integration into the chromosome the downstream genes are expressed. In this

study, we used the LIC methodology to generate C-terminal SPA tag (sequential peptide affinity)

translational fusions with genes of interest, through a pMUTIN-SPALIC integrative plasmid

containing a LIC site generated synthetically.


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Figure 66. LIC integration into B. subtilis. (A) The insert, corresponding to the last 400bp of the gene of

interested, was amplified using the forward and reverse primers P1 and P2. These primers contain specific

sequences homologous to the pMUTIN-SPA vector allowing the hybridization between the insert and the vector.

After mixed the insert with the vector, the sample was transformed into E.coli MC1061 strain, a step that allows

the amplification of the plasmid. (B) Thereafter, the final product plasmid was transformed into B. subtilis and

inserted into the chromosome by simple cross-over. Then a fragment containing the gene of interest fused to the

SPA tag was amplified from the genomic DNA of the final strain and sequenced.

A

B


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4.1.12 DNA sequencing

Confirmation of DNA sequences was outsourced to GATC Biothec. The sequencing

chromatograms were analysed and the sequences were confirmed using the Geneious software

alignment features, using as background the BSB1 complete genome.

4.1.13 DNA transformation

4.1.13.1 Transformation of Escherichia coli (Heat shock)

The transformation of DNA into E. coli MC1061, BL21 or ER2566 chimiocompetent strains

were performed by heat shock. 20-50µl of the frozen competent stocked cells were mixed with the

LIC sample, ligation mix or 1-5µl of 10pg to 100 ng/µl of plasmid DNA and incubated on ice during

20 minutes. The mixture was heat shocked for 45s at 42˚C and then put on ice again for 2 min. 500 µL

of fresh LB were added to the cells and incubated at 37˚C for 45 min. Finally, the cells were harvested

by centrifugation (1 min at 12000 xg), resuspended in 100 µL of fresh LB and plated on selective agar

medium.

4.1.13.2 Transformation of Bacillus subtilis using natural competence

Transformation of B. subtilis cells was performed using a quick procedure protocol which is

based on the natural competence of vegetative cells. A fresh plate was prepared by spreading the stock

culture on plate, supplemented with the appropriate selection markers, and incubated overnight at

37˚C. After overnight incubation, a single colony was used to inoculate 2 ml of MC complemented

medium (see appendix,section 6.1) and the culture was grown for 5h, at 37°C in a shaking incubator

(200 rpm). Then 400µl of culture were mixed with 10µl of plasmid DNA or 1µl of chromosomal DNA

and grew for additional 2 hours at 37°C with agitation (200rpm). The culture was plated on LB plates

supplemented with the appropriate antibiotic and incubated overnight at 37°C.

4.2 Bacterial strains and growth conditions

E. coli and B. subtilis strains used in this work are listed in Table S2 and Table S3

respectively. B. subtilis strains used were derived from the BSB1 reference strain, which is a

tryptophan-prototrophic (trp+) derivative of B. subtilis 168-trpC2 strain (Nicolas et al. 2012).

Nutrient Luria-Bertani medium (LB) was used for general-purpose growth of E. coli and B.

subtilis. For B. subtilis DNA transformation, MC media was used (described at the transformation of

B. subtilis section). For specific experiments we used CH defined medium, M9 or Belitsky minimal

media. In general, the bacterial growth was at 37 °C with orbital shaking at 200 rpm, unless otherwise

stated. Growth supplements (Table S 1) and antibiotics (Table S 2) were added to the media when

appropriate. Growth was monitored by measuring optical density at 600 nm (OD600nm) in a

spectrophotometer. For long-term storage of bacterial strains, LB broth cultures, after overnight


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growth, were supplemented with 30 % (v/v) glycerol and stored at -80 °C in cryovials. Growth media

and buffers are described below in the Appendix.

4.3 Strain construction

4.3.1 SPA tagged strains construction

The utilization of a SPA tag allows the purification of proteins in their native conditions. In

order to study the possible protein-protein interactions in between the proteins involved in the Sec

machinery, we have constructed translational fusions of a SPA-tag DNA sequence to the to the 3’ end

of the gene of interest. The cloning of a selected group of proteins involved in the Sec machinery

(secDF, secG, spoIIIJ, bdbB, bdbC, sipT, sipS, sppA, yteJ and prsA), into the BSB1 background, were

carried out according the LIC system described in section 4.1.11. The fragments contained ≥400bp of

the end of the gene of interest, excluding the stop codon, and were amplified from chromosomal DNA

using the appropriate primers listed in Table 2. The primers had a designed LIC tail sequence

homologous to the LIC sequence present on the pMUTIN-SPA LIC plasmid. The vector and the insert

were processed as described on the section 4.1.11 and Figure 66. After the annealing, of the vector

and insert, at room temperature, the mixture was transformed into E.coli MC1061 and the

transformants were selected in 100 µg.ml-1 ampicillin. The final product plasmids were extracted

from E. coli and used to transform BSB1 strain. B. subtilis transformants were selected on LB agar

plates supplemented with 0.6 µg.ml-1 Erythromycin. The 400 bp homology with the targeted gene

allows the plasmid to insert into the chromosome by a single crossover in B.subtilis. To select the

positive clones, the colonies that grew in the presence of 0.6 µg.ml-1 Erythromycin were re-streaked

into 30 µg.ml-1 and grow at 37°C for overnight. The final strains harbouring a C-terminal SPA fusion

under the control of the gene native promotor are listed at Table S 4.

To obtain N-terminal SPA tagged strains, the whole sequences of ORF corresponding to the

genes of interest were amplified by PCR from the BSB1 chromosomal DNA, including the start codon

and stop codon. For this, we used a specific pair of primers for each gene, carrying NotI and XhoI

restriction sites at their 5’ and 3’ ends, respectively. The resulting PCR product was digested with NotI

and XhoI restriction enzymes and ligated into pSPANter plasmid, previously digested with the same

enzymes. E. coli MC1061 cells were transformed with the ligation mixtures and, then, single colonies

were selected on LB-agar plates supplemented with 100 µg.ml-1 ampicillin. The plasmids harbouring

the N-terminal SPA construction were transformed into BSB1 and integrated into the amyE locus by

double crossing over. This integration gives origin to the SPA-tagged protein of interest expressing

strains, under the control of the xylose inducible promoter Pxyl. The positive clones, were selected in

100 µg.ml-1 spectinomycin, confirmed by PCR from the amyE flanking region, tested by the inability

of the strain to produce amylase (starch-iodine test) and sequenced. All the strains constructed are


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listed in Table S2 and S3 of appendix. The final strains harboring a SPA fusion were confirmed by

PCR and sequenced.

The starch-iodine test is used to check the presence of starch, when present the starch reacts

with the iodine crystals and turns into an intense blue/dark color (Dhar 1923). As the amyE gene

encodes for an amylase, this test allows a quick selection of the positive clones. The insertion of the

gene of interest disrupts the amyE gene, consequently the starch is cannot be not degraded, so when

the clones are streaked into a plate containing starch the positive ones turn dark in the presence of

iodine.

Table 2 Oligonucleotides used for LIC constructions

(Sequences required for LIC are underlined)

Sequence

amplified

Primer name Primer sequence

secDF secDF_P1 GGGTTCCTGGCGCGAGCTTGCTATAGCTTCTATTGG

secDF_P2 TTGGGCTGGCGCGAGC

TTGCGCCGAATCTTTTTTCAG

secDF_seq ACGGATCTGATCCAAATGTCAGC

secG secG_P1 GGGTTCCTGGCGCGAGCCACGCAGTTTTGATTACC

secG_P2 TTGGGCTGGCGCGAGC TAGGATATAAGCAAGCGC

secG_seq TCTGATACAGCGTGACATGC

spoIIIJ SpoIIIJ_P1 GGGTTCCTGGCGCGAGC

AGCTTCAACAGGAAACAATGGC

SpoIIIJ_P2 TTGGGCTGGCGCGAGC CTTTTTCTTTCCTCCGGC

SpoIIIJ_seq AGTTCGAAAGCGATGCAGGC

bdbB BdbB_P1 GGGTTCCTGGCGCGAGC

TGGCACAATGGCTAGTTTATTC

BdbB_P2 TTGGGCTGGCGCGAGC ATTATATACATGTTGATTTTG

BdbB_seq TCTTGCCAACTCACTCCAC

bdbC BdbC_P1 GGGTTCCTGGCGCGAGCTATGCTTCCTGGGTTGTG


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BdbC_P2 TTGGGCTGGCGCGAGCTTCAGATTTTTCGCCTTTCAG

C

BdbC_seq TGCGGATTATGATGAAATC

sipS SipS_P1 GGGTTCCTGGCGCGAGC

TGACTCTATGTATCCTACAC

SipS_P2 TTGGGCTGGCGCGAGC ATTTGTTTTGCGCATTTCG

SipS_seq AGCAATTGTGATTGCTGTCG

sipT SipT_P1 GGGTTCCTGGCGCGAGC

CACATTACATGACGGAGAAAGG

SipT_P2 TTGGGCTGGCGCGAGC TTTTGTTTGACGCATTTCG

SipT_seq TCATCGCTGTTCTGCTGGC

sppA SppA_P1 GGGTTCCTGGCGCGAGCCCATAAGGACATTATGTCTC

SppA_P2 TTGGGCTGGCGCGAGCCTTCGCATAGAGATACATC

SppA_seq TCGGCGTCATTATGGAAAGC

yteJ YteJ_P1 GGGTTCCTGGCGCGAGC TTCCTGCTTGATTGGCTC

YteJ_P2 TTGGGCTGGCGCGAGC

TTTGCGGTATAATTTTTCATGC

YteJ_seq AGAGTTAGAGCGCAATGACATC

prsA PrsA_P1 GGGTTCCTGGCGCGAGC

AGGCGAGAAGTTTGAAGACC

PrsA_P2 TTGGGCTGGCGCGAGC TTTAGAATTGCTTGAAGATG

PrsA_seq AAGGCAAAATCCGTGCAAGC

4.3.2 Deletion mutants

The deletion of sppA, yteJ and the sppA yteJ operon at their native locus in the BSB1 strain

was performed by sequence replacement with a chloramphenicol resistance cassette (CmR) expressed

under the control of the native promoter. Therefore, PCR fragments were obtained by amplifying the


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upstream and downstream regions, approximately 1 Kb each, of the gene sequence, from the BSB1

genomic DNA in two sequential PCR steps using the appropriate pair of primers, KO-P1 plus KO-P2

and KO-P5 plus KO-P6 respectively, listed in Table S1. Then a resistance cassette of chloramphenicol

(CmR), approximately 1 Kb, was amplified from pUC19::chloramph plasmid using the primers KO-P3

and KO-P4. The primers P2, P3, P4 and P5, of each construction, have an extension containing an

overlap sequence between the upstream, chloramphenicol and downstream fragments. These three

amplified products were then purified and joined by an overlap PCR reaction. The final PCR product

was purified from agarose gel, directly transformed into BSB1 and plated on LB agar supplemented

with 5 µg.ml-1 chloramphenicol. The deletion of the genes of interest from the native locus was

confirmed by PCR and the final constructs were sequenced, resulting in GHBs023, GHBs025 and

GHBs028 strains. Primers sequences and strains are listed inTable S1 and Table S2 of appendix.

4.3.3 SppA, YteJ and SppA-YteJ overexpressing strains

With the goal of re-insert sppA and yteJ into the deletion mutants, under the control of

inducible promotor, to observe if we could recover the effect on lantibiotics resistance caused by their

deletion we have constructed strains overexpressing sppA and yteJ. To obtain strains overexpressing

SppA or YteJ, or both, we inserted the gene sequences fused to the Phyperspank promoter in the BSB1

chromosome at the amyE locus. For this, the entire gene sequences of sppA and yteJ and the sppA-yteJ

operon, including the RBS sequence, were amplified by PCR from BSB1 genomic DNA , and fused to

the Phyperspank promoter sequence. This promoter is Isopropyl-β-D-thiogalactopyranoside (IPTG)

inducible and lacI-repressible promoter (Overkamp et al. 2013),. For this, the RBS+genes sequences

were amplified by PCR using a specific pair of primers, the Hspank_P1 and Hspank_P2 of each gene,

carrying the HindIII and SalI restriction sites. The resulting PCR products were digested with HindIII

and SalI fast restriction enzymes and ligated into pDR111 plasmid, previously digested with the same

enzymes. HindIII cuts into the sppA sequence, consequently the constructions involving this gene

were done with a blunt 5’ end and a sticky 3’ end created by SalI digestion. The vector and the

fragments were ligated and transformed into E. coli MC1061 competent cells. After the selection of

transformants on 100 µg.ml-1 ampicillin LB-agar plates, the plasmids were extracted, purified and

transformed into BSB1. The positive clones, harboring an integration of the Phyperspank+gene of

interest at the amyE locus, were selected on 100 µg.ml-1 spectinomycin LB-agar plates, confirmed by

PCR using primers in the amyE flanking regions, tested for the inability of the strains to produce

amylase (starch-iodine reaction) and sequenced. With the objective of complementing the deletion

mutants by reintroducing the deleted gene at the ectopic amyE locus, under the control of the inducible

Phyperspank promoter, the final plasmids were also transformed into deletion mutant strains (GHBs023,

GHBs025 and GHBs028). Table S2 of appendix.


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4.3.4 Construction of the GFP Fusions

Two types of superfolder green fluorescent protein (sfGFP) (Pédelacq et al. 2006) fusions

were constructed during this study: transcriptional fusions with the native σW and σA-dependent

promoters, of sppA yteJ operon and translation fusions of SppA and YteJ.

The expression of the sppA-yteJ operon is under the control of two adjacent promoters,

dependent of σW and σA. To study the sppa-yteJ native promoters activity we constructed two strains

harboring a fusion of one and the two promoters with the DNA sequence of the sfgfp introduced at the

ectopic locus amyE.. First, a fragment of 100bp upstream of sppA, containing the natural RBS and the

σW promoter sequence, and a 185bp fragment, including the natural RBS, σW and σA promoter

sequences (Nicolas et al. 2012), were amplified from BSB1 genomic DNA by PCR, and then ligated

upstream of the pSG13 plasmid-borne sequence of the fluorescent protein (Guiziou et al. 2016) by

Gibson assembly. The final mix was transformed into E. coli MC1061 competent cells and plated onto

100 µg.ml-1 ampicillin LB-agar. The plasmid was extracted from positive E. coli clones purified and,

after transformation of B. subtilis BSB1 competent cells, was integrated into the amyE locus. The

positive clones were selected onto 100 µg.ml-1 spectinomycin LB-agar plates, the presence of the

promoter-sfGFP sequence fusions were confirmed by PCR, the clones were also tested by starch-

iodine reaction and, finally, sequenced. The final strains were tested by LCA, as described on section

4.5.

To study SppA and YteJ localization, N-terminal tagged sfGFP-SppA and C-terminal tagged

YteJ-GFP proteins were conditionally expressed from the IPTG inducible Phyper-spank promoter, at the

ectopic amyE locus of B. subtilis chromosome. The appropriate primers, one carrying a HindIII or SalI

restriction site and the second carrying an extension with a linker, were used to PCR amplify the sppA

and yteJ genes from BSB1 chromosomal DNA and the sfgfp gene from the pSG13 plasmid. These

fragments were joined by PCR, digested by HindIII or SalI and ligated into the pDR111 plasmid,

previously digested with the same enzymes. E. coli MC1061 cells were transformed by the ligation

mixtures and transformants were selected onto 100 µg.ml-1 ampicillin LB-agar. The plasmid was

extracted from positive E. coli clones and purified. The sfgfp-sppA and yteJ-sfgfp constructs were then

integrated at the amyE locus of BSB1 competent cells by transformation, and transformant clones

were selected using 100 µg.ml-1 spectinomycin LB-agar plates. The presence of the integrated sfGFP

fusions were verified by PCR amplification from the amyE flanking regions and by the inability of the

strain to produce amylase. The final constructs were sequenced.

4.3.5 His- tag constructions

To obtain E. coli strains overexpressing N-terminal 6xHis tagged proteins construct, the genes

of interest were amplified by PCR from the BSB1 chromosomal DNA, excluding the start codon. For

this, we used a specific pair of primers for each gene, carrying NdeI and XhoI restriction sites (


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Table S 3). The resulting PCR product was digested with NotI and XhoI fast restriction

enzymes and ligated into pJ411_6His, previously digested with the same enzymes, a plasmid

harboring the 6XHis-tag sequence under the control of the PT7 promoter. E. coli MC1061 cells were

transformed by the ligation mixtures and transformants were selected onto 30 µg.ml-1 kanamycin LB-

agar plates. The plasmids were extracted by minipprep and the presence of the right inserts were

confirmed by PCR and sequenced. Thereafter, the final plasmids were used to transform into E.coli

2566 overexpressing strains. The final strains obtained, from GHEc 054 up to GHEc 058, is listed on

Table S 4.

4.4 Fluorescence Microscopy

B. subtilis strains were grown overnight in LB at 37 ºC with the appropriate antibiotic and, the

next day, were diluted (1:200) in 20 ml of fresh LB supplemented with 0.05 mM IPTG and grown

until OD600nm 0.3-0.5 - exponential growth phase - and until OD600nm 1-2 - early stationary growth

phase. Thereafter at regulated intervals samples were taken and cells were observed by fluorescence

microscopy. For that, 2 µl of cell culture were placed onto a thin film of 1.2% agarose. The cell

membranes were stained with FM4-64 (Molecular probes) and nucleoids were stained with DAPI.

Fluorescence microscopy was performed using a Leica DMRA2, controlled by Metamorph software

(Molecular devices). Processing of fluorescence images was performed using Metamorph and ImageJ

software.

4.5 Live-cell array (LCA)

The Live Cell Array is a method (Botella et al. 2010), which allows the high throughput

acquisition of time-course profiles of bacterial cellular growth and gene expression. It measures both

the optical density for recording cell growth and the fluorescence emission of the fluorescent reporter

protein, usually the GFP, expressed in specifically modified strains of interest. The B. subtilis strains

were grown overnight in 5ml of LB medium at 37°C and then diluted 20-fold in 100µL/well of fresh

LB medium (pre-culture) into 96-well microtiter plate. The LB pre-cultures were grown until an

OD600nm of 0.3-0.4 and then diluted 400-fold into a fresh medium, LB or CH defined medium. OD600nm

and fluorescence (excitation 485, emission 528) were measured at time intervals of 10 minutes. Only

the 60 central wells of the 96-well microtiter plates were inoculated since the wells around the edges

exhibited evaporation higher than 5 % over 20 hours; therefore these peripheral wells were only filled

in with sterile medium.

4.6 Bacteria survival test

The BSB1 and deletion mutant strains, described in section 2.1.2, were grown overnight in LB

medium, supplemented with the appropriate antibiotic, at 37°C. On the following day, the overnight


142

cultures were diluted 200-fold into fresh LB or M9 medium and then the cultures grew for 48h at 37

°C while shaking. Samples of 100µl were taken after 18h, 24h and 48h. Serial dilutionswere done and

the last two dilutions, 10-6 and 10-7 fold, were plated onto LB agar supplemented with the appropriate

antibiotic. The number of colonies, corresponding to the number of cells that survived, were counted

after overnight incubation at 37°C.

To test if the WT cells could complement or help the growth of the mutants, the WT cells and

deletion mutants were mixed and grew for 48h at 37 °C while shaking. The overnight cultures were

diluted 200-fold into fresh LB and grown at 37°C until reach the exponential phase, then the cultures

were diluted 1000-fold and mixed 1:1 (50% WT culture+50% Deletion mutant). The mixed cultures

and the controls grew for 48h at 37 °C while shaking. From this point the remaining steps were done

as described above.

4.7 Swimming and swarming capacity

The surfaces of agar concentrations above 1% prohibit swarming and swimming of many

bacterial species, consequently, the humidity of the media is crucial for conducting these experiments.

Therefore, it is necessary to define a rigorous protocol in order to control the water content of the

medium, we tested the ability of the bacteria swim at 0.3% of agar and the capacity of swarm at 0.7%

(Kearns 2010). All the plates were consistently prepared, 25 ml of the LB media were poured when

the agar is approximately 50°C, let to solidify and to dry open for 15 minutes - in a laminar flow

chamber. The cells were harvested (100µl) from overnight cultures or at mid exponential phase

(OD600nm ~0.5), centrifuged at 2000rpm to avoid breaking the flagella, resuspended in PBS buffer and

spotted (5µl) in the middle of the plate. Thereafter, the halo created by the cell movement were

measured at time intervals of 1h during 6h.

4.8 Biofilm formation

B. subtilis strains in study were grown in LB shaking at 37°C, supplemented with the

appropriate antibiotic, to an OD600nm of 1,0. To observe the architectural differences between biofilm

from distinct strains, 10 µL cultures were then spotted on a dried MSgg plate (1.5% agar) and

incubated for 70 hours at 30 °C. Colonies were measured and photographed.

4.9 Protein analysis

4.9.1 Sodium Dodecyl Sulphate Poly-Acrylamide Gel Electrophoresis (SDS

PAGE gels)

All the proteins purified in this project were analysed by 10% or 12.5% SDS-PAGE gels.

Samples were mixed with protein loading buffer (0.5 M Tris-HCl pH = 6.8, 1 M β-mercaptoethanol,

20 % (v/v) SDS, 0.2% (v/v) bromophenol blue, 12.5 % (v/v) glycerol), usually incubated 5 min at 95


143

°C and electrophoresis was performed in Laemlli 1X buffer (25 mM Tris, 192 mM glycine, 0.1%

SDS) at 200 V. After staining with Coomassie blue staining solution (0.1% Coomassie Brilliant Blue

R-250, 50% ethanol and 10% glacial acetic acid) protein sizes were estimated relative to a protein

marker ladder.

4.9.2 Blue Native PAGE

For Blue-Native electrophoresis, the strains of interest were pre-cultured overnight in LB,

supplemented with antibiotics if needed, at 37°C. The following day the culture was diluted 1000-fold

(to an OD600nm of 0.001) into 1-2 liters of LB medium, supplemented if needed. When the OD600nm

reached 0.5 - 1, cells were harvested by centrifugation at 6000 rpm for 10 min at 4°C. The pellet was

washed with 40 ml of cold resuspension buffer (buffer R: 10 mM Tris-Cl pH 7.5, 150 mM NaCl),

centrifuged at 4500 rpm for 10 min at 4°C and instantly frozen in liquid nitrogen. The cells were

resuspended in 20 ml of buffer R supplemented with 1 mg/ml of lysozyme, 1 mM EDTA and 5 µl of

Benzonase (Invitrogen), and incubated in a water bath at 37 °C for 15-30 min. To remove the cell

debris the samples were centrifuged at 4500rpm for 30min at 4°C and the pellet was discarded.

Membrane and cytosolic fractions were then separated by ultracentrifugation at 100,000 g for 1 h at

4°C. The membrane protein samples were solubilized into 1% n-Dodecyl β-D-maltoside (DDM), were

mixed with NativePAGE sample buffer and with NativePAGE 5% G-250 Sample Additive

(Invitrogen), then incubated for 1h on ice. The samples were separated by electrophoresis on

NativePAGE 3%–12% Bis-Tris gels (Invitrogen) with NativePAGE running buffer and NativePAGE

Dark Blue Cathode Buffer (Invitrogen) for 20 to 30 min at 150 V, substituted with NativePAGE Light

Blue Cathode Buffer (Invitrogen) for 60 min at 150V. The electrophoresis tank was maintained on ice

throughout the procedure. The gel was then used for Western blot analysis or stained by Blue Safe

Protein Stain. This staining allows the analysis by nanoscale liquid chromatography coupled to tandem

mass spectrometry (nanoLC-MS/MS, Plateforme PAPPSO, INRA centre de Jouy en Josas, or Dr

Becher’s lab, Greifswald University).

4.9.3 Western blot analysis

To analyze if the tagged proteins were expressed, Western blots were performed using the

appropriate primary and secondary antibodies for each specific tag. In the case of FLAG-tag the

Monoclonal ANTI-FLAG M2 antibody (F3165 Sigma) as a primary antibody and Anti-Mouse IgG

(A2304 Sigma) as a secondary antibody. For the AmyM the primary antibody, we used, was raised

during my UCB secondments, this antibody was raised in rabbits, as such we used Anti-Rabbit IgG

(A0545 Sigma) (Appendix section 6.2) Equal amounts of protein, from each sample, were heated to

100 ºC for 5 minutes, loaded onto a 10% SDS-PAGE gel and separated at 200V. Proteins were then

transferred to a Hybond-P Polyvinylidene fluoride (PVDF) membrane (GE Healthcare) using a

semidry transfer cell (Bio-Rad) according to standard Western blotting techniques(Burnette 1982).


144

The detection was performed, after a last wash step, using Clarity™ Western ECL Blotting detection

substrates (Biorad), according to the manufacturers guidelines, through the ChemiDoc™ XRS+

System and Image lab imaging software.

4.9.4 Secreted proteins analysis

To check if a protein complex is involved in protein secretion we analyzed the if extracellular

amount of a highly secreted reporter protein, as the case of AmyM, is affected. For that purpose, the

B.subtilis strains of interest were naturally transformed with a multi copy plasmid containing the gene

encoding AmyM (pCS73) and an empty plasmid (pCS74) to use as a control, all carrying a neomycin

(Neo) resistance cassette. Dr. Van Rij provided these plasmids from DSM company, our collaborator

from the Protein Factory project. The final strains, carrying the pCS73 and pCS74, were pre-cultured

overnight at 37°C in LB medium supplemented with 10µg/ml Neo and other antibiotic markers

correspondent to the background strains, if needed. The following day the culture was diluted 1000-

fold (to an OD600nm of 0.001) into 50ml of LB medium with the same supplements. Grown at 37°C

until the OD600nm reached 0.5-1 and the cells were harvested by centrifugation at 4500 rpm for 15 min

at 4°C. The cell pellet was discarded and the culture media was filtered using a 0.2µm filter, removing

possible remaining cells. The clear culture media was precipitated overnight at 4°C by 10%

trichloroacetic acid (TCA). After centrifugation at 15000 rpm for 30 min, the supernatant was

discarded and the protein pellet was washed two times with 200 µl cold acetone, repeating the

centrifuge steps each time. The protein pellets were then dissolved by the addition of 3µl of 10 mM

Tris-HCl pH 8, 8 M urea.

4.9.5 Sequential Peptide Affinity (SPA) purification

The sequential peptide affinity (SPA) technique has been developed, from the TAP-tag

technique, to study protein-protein interactions (Rigaut et al. 1999; Zeghouf et al. 2004). The

translational fusion of the SPA-tag, composed of three FLAG sequences (3×FLAG) and the

calmodulin binding peptide (CBP) separated by a TEV protease cleavage site, to a protein of interest

allows its purification together with the protein(s) to which it interacts under native conditions, i.e.

without adding any cross-linking agents ( Delumeau et al. 2011).

The strains expressing SPA-tagged proteins were pre-cultured overnight in the presence of

erythromycin at 37°C. The following day the culture was diluted 1000-fold (to an OD600nm of 0.001)

into 2 liters of LB medium supplemented with 0.6 µg/ml erythromycin and 0.5 mM IPTG. When the

OD600nm reached 0.5 - 1, cells were harvested by centrifugation at 6000 rpm for 10 min at 4°C. The

pellet was washed with 40 ml of cold resuspension buffer (buffer R: 10 mM Tris-Cl pH 7.5, 150 mM

NaCl), centrifuged at 4500 rpm for 10 min at 4°C and instantly frozen in liquid nitrogen. The cells

were resuspended in 20 ml of buffer R supplemented with 1 mg/ml of lysozyme, 1 mM EDTA and 5


145

µl of Benzonase (Invitrogen), and incubated in a water bath at 37 °C for 15-30 min. To remove the cell

debris the samples were centrifuged at 4500rpm for 30min at 4°C and the pellet was discarded.

Membrane and cytosolic fractions were then separated by ultracentrifugation at 100,000 g for 1 h at

4°C. The membrane pellet was solubilized using buffer R supplemented with 1% DDM. Identical

protein amounts of the membrane and soluble protein fractions were loaded on 10% SDS-

polyacrylamide gels for Coomassie staining and Western blotting using anti-FLAG antibodies.

In addition, protein complexes from the membrane fraction were pulled down using a SPA

purification method (Charbonnier et al. 2017). The solubilized membrane fraction, 2 mM CaCl22 and

1%DDM, was mixed with 200 µl of pre-equilibrated calmodulin resin (CaM) in 2 ml tubes, which

were placed on a rotating wheel for a minimum of 2 h at 4°C. The resin was transferred in a column,

the eluate was discarded and the CaM resin was washed with 2 ml of buffer R supplemented with 1%

DDM and 0.1 mM CaCl2. The proteins remaining bound to the CaM resin were eluted with a total

volume of 0.75 ml of buffer R supplemented with 1% DDM and 3 mM EGTA. The eluate was

collected and mixed with 200 µl of anti-FLAG resin. After its incubation under gentle agitation on the

rotating wheel for, at least two hours, the eluate from the anti-FLAG resin was discarded, the resin was

washed with 2 ml of buffer R supplemented with 1% DDM. The proteins remaining bound to the anti-

FLAG resin were eluted with a total volume of 1ml of 0.1M glycine pH 3.

Proteins from the eluate were precipitated overnight at 4°C by 10% trichloroacetic acid

(TCA). After centrifugation at 15000 rpm for 30 min, the supernatant was discarded and the protein

pellet was washed two times with 200 µl cold acetone, repeating the centrifuge steps each time. The

protein pellets were then dissolved by the addition of 3µl of 10 mM Tris-HCl pH 8, 8 M urea. The

volume was then extended up to 25µl with 10 mM Tris-HCl pH 8, decreasing the urea concentration to

1 M. Then to analyze if the SPA tagged protein was captured, with possible interacting partners, the

protein concentrations of the samples were measured by the Bradford’s method and the same amount

of protein was loaded and the samples were separated by native polyacrylamide gel electrophoresis

(PAGE), which was coomassie stained after. These samples were analyzed by a second technique,

nanoscale liquid chromatography coupled to tandem mass spectrometry (nano LC-MS/MS). For this

the samples were equal loaded into SDS-Page gel, as before, but the samples just run 1 cm into the

resolving part of the gel. The gel was stained by Bio-Safe Coomassie G-250 stain solution and the

small pieces of the gel containing the samples were cut and sent to be treated and analyzed by mass

spectrometry.


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Figure 67. Sequential peptide affinity purification. (A) The SPA-tag is composed of three FLAG sequences

(3×FLAG) and the calmodulin binding peptide (CBP) separated by a TEV protease cleavage site. (B) Schematic

representation of the double step SPA purification. This technique allows the purification of protein complexes

under native conditions.

4.9.6 His-tag purification

The E. coli strains harboring a translational 6XHis-tag fusion were pre-cultured overnight in

the presence of 30 µg.ml-1 kanamycin at 37°C. The following day the culture was diluted 1000-fold (to

an OD600nm of 0.001) into 2 liters of LB medium supplemented with 30 µg.ml-1 kanamycin and grown

until an OD600nm of 0.8 at 37°C. Thereafter the expression was induced by the addition of 250 µM

IPTG for overnight at 16°C. Cells were harvested by centrifugation 4500 rpm for 10 min, and the

pellet was resuspended in 40 ml of 20mM Tris-HCl pH7,5 and 250mM NaCl (BufferR2) and stored at

-20°C. Cells were lysed by sonication and centrifuged at 4500rpm for 30 min to remove the cell

debris. The membrane and cytosolic fractions were separated by ultracentrifugation at 100,000 g for 1

h at 4°C. The membrane fraction was solubilized using buffer R2 supplemented with 1% n-Dodecyl β-

D-maltoside (DDM) for overnight at 4°C. The sample was then loaded onto a Ni2+ affinity column

(Ni-NTA agarose, Qiagen) pre-equilibrated in the same buffer. The Ni-NTA columns were washed

with 50ml of 20mM Tris-HCl pH7,5, 250mM NaCl and 0.01% DDM (Buffer R3) supplemented with

5mM imidazole, and then with 20 ml Buffer R3 suplemented with 20mM imidazole. After washing,

the proteins bound to the Ni-NTA resin were eluted, sequentially, with 1ml of Buffer R3 supplemented

with 50 mM imidazole and then 4ml of the same buffer with 200mM imidazole. All the eluted samples

A

A

A

A

A

A

A

B

B

B

B

Fig

ure

1|

Seq

ue

nti

al

pe


147

were analyzed by SDS-PAGE. The samples containing the highest concentration of the His-taged

protein were joined and concentrated by Amicon Ultra-15 Centrifugal Filters, mixed with 50%

glycerol and stored at -20°C.

4.9.7 Double tag purification

With the objective of increasing the purity of the membrane protein complex, a E. coli stain

harboring SppA with a N-terminal His tag and YteJ with a C-terminal Flag ta ( 6xHis-SppA/YteJ-Tev-

3xFlag), a double tagged SppA-YteJ complex, was constructed as described above on section 4.3.5.

The membrane fraction was obtained and solubilized as described above. The sample was mixed with

200 µl of anti-FLAG resin and incubated under gentle agitation on the rotating wheel. The eluate from

the anti-FLAG resin was discarded and the resin was washed with 2 ml of buffer R supplemented with

1% DDM. The resin was resuspended in 200 ml of TeV buffer (50 mM Tris-HCl pH 8, 150 mM NaCl,

0.5mM EDTA, 1 mM DTT) with 50units of Ac-TeV protease. After incubation at 4°C for overnight,

the eluate was collected, diluted up to 2ml (4-fold dilution) and mixed with 200µl Ni-NTA pre-

equilibrated agarose resin. The Ni-NTA columns were washed with 50ml 20mM Tris-HCl pH7,5,

250mM NaCl and 0.01% DDM (Buffer R3) supplemented with 5mM imidazole ,and then with 20 ml

Buffer R3 supplemented with 20mM imidazole. After washing the proteins bounded to the Ni-NTA

resin were eluted, sequentially, with 1ml of Buffer R3 supplemented with 50 mM imidazole and then

4ml of the same buffer with 200mM imidazole. The eluted proteins were concentrated by Amicon

Ultra-15 Centrifugal Filters, mixed with 50% glycerol and storage at -20°C.


148

149

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177

APPENDIX

Chapter 6. APPENDIX

179

6 APPENDIX

6.1 Contents, sterilization and storage of required

solutions

Luria Bertani (LB) medium- For 1L of media join 10 g Bacto-Tryptone (Becton Dickinson

and Co); 10 g NaCl (Merck); 5 g Yeast extract (Becton Dickinson and Co) and nanopure H2O up to

1L. Medium can be autoclaved at 121°C for 20 minutes and stored at RT.

Luria Bertani (LB) agar medium- For 1L of media join 10 g Bacto-Tryptone (Becton

Dickinson and Co); 10 g NaCl (Merck); 5 g Yeast extract (Becton Dickinson and Co); 15 g Agar

(Invitogen) and nanopure H2O up to 1L. Medium can be autoclaved at 121°C for 20 minutes and

stored at RT.

M9 minimal medium- For 1L of media add the following solutions (that were previously

sterilized separately): 1 ml 100 mM CaCl2; 10 ml trace saltsA

100x stock; 1 ml 1 M MgSO4; 1 ml 50

mM FeCl3/100 mM C6H8O7; 6 ml of glucose 50% (w/v) stock solution and 200 ml M9B

5x stock.

Trace salts 100x stock : 100 mg MnCl2.4H2O (Merck), 170 mg ZnCl2 (Sigma); 43 mg

CuCl2.2H2O (Merck); 60 mg CoCl2.6H2O (Merck); 60 mg Na2MoO4.2H2O (Aldrich), per liter.

Contrary to what is described by Harwood and Cutting (1) MgSO4, CaCl2 are added as separate

solutions to prevent precipitation problems. FeCl3 is also added as a separate solution as this

compound is unstable. Filter-sterilize the solution and store in the dark at RT.

M9 5x stock: 42.5 g Na2HPO4.2H2O(Merck); 15 g KH2PO4 (Merck); 5.0 g NH4Cl (Merck);

2.5 g NaCl (Merck). Adjust to pH 7.0 using 4M NaOH. Solution is autoclavable at 121°C for 20

minutes and stored at RT.

Belitsky-Minimal Medium- For 1L of media join 2 g (NH4)2SO4; 2 g MgSO4 * 7H2O; 2g

KCl, 2g Na3-Citrate * H2O and 50 ml 1M Tris-HCl pH 7.5. Add nanopure H2O up to 1L and adjust pH

to 7.5. Then this media needs to be supplemented with 5 ml 0.2M KH2PO4; 9 ml 0,5 M Glutamic

Acid; 10 ml 20% Glucose , 2 ml 1M CaCl2; 2 ml 0,0005M FeSO4 and 0,4 ml 0,025M MnSO4.

Completed Competence Medium (MC completed)- For 100ml of solution: 14g

K2HPO4*3H2O, 5.2g KH2PO4, 20g glucose, 8.8g C6H5Na3O7, 2.2g C6H5O7FeNH4, 1g casein

hydrolysate, 2g C5H8KNO4*H2O and 3.35mM MgSO4.

MSgg medium- it is composed of 5mM K3PO4 (pH7),100mM MOPS (pH7), 2mM MgCl2,

700μM CaCl2, 50μM MnCl2, 5μM FeCl3, 1μM ZnCl2, 2μM thiamine, 0.5% glycerol, 0.5% glutamate,

50 μg ml–1 tryptophan and 50 μg ml–1 phenylalanine.

Chapter 6. APPENDIX

180

Subtilin preparation- The subtilin is produced by the B.subtilis ATCC6633 strain, which was

supplied by Jan Maarten van Dijl group. To extract it, this strain was grown overnight in LB medium

at 37°C. The supernatant was collected by centrifugation at 1400 rpm for 10 min, the pellet was

discarded. Aliquots of 25 ml of supernatant were heated at 80°C for 10 min and filtered. These

samples were freeze with liquid nitrogen and then freeze dry overnight. Initial experiments were done

with 30% m/v subtilin. However, to avoid the effect of volume in the 96 well plate culture we

concentrated 10X times the stock solution. The sample powder was weighed and concentrated 10

times by resuspensing in water, by other words the stock powder corresponding to 25 ml of liquid

sample was ressuspended into 2.5 ml. This 10X stock solution was then used to induce the strains at

30% v/v or other concentrations when needed for testing. The powder sample was kept at -20°C.

Table S 1. Stock and working solutions of growth supplements

Supplement Solute Stock concentration Usual working

concentration

IPTG dH2O 1 M 0.5 mM

Xylose dH2O 20% 1%

Table S 2. Stock and working solutions of antibiotics.

Antibiotic Solute Stock concentration

(mg.ml-1)

Working

concentration

(µg.ml-1) E.coli

Working

concentration

(µg.ml-1) B.subtilis

Ampicillin (Amp) dH2O 50 100 -

Spectinomycin

(Spec) dH2O 50 - 100

Neomycin (Neo) dH2O 50 - 10

Erythromycin

(Erm) 100% ethanol (v/v) 6 - 0.6 or 30

Chloramphenicol

(Cm) 50% ethanol (v/v) 10 - 5

Kanamycin (Kan) dH2O 50 30 -

Chapter 6. APPENDIX

181

Table S 3 . List oligonucleotides

Construction Primer name Sequence (5’-> 3’)

C-terminal SPA tag

fusion

SecDF_P1 GGGTTCCTGGCGCGAGCTTGCTATAGCTTCTATT

GG

SecDF_P2 TTGGGCTGGCGCGAGC

TTGCGCCGAATCTTTTTTCAG

SecDF_seq ACGGATCTGATCCAAATGTCAGC

SecG_P1 GGGTTCCTGGCGCGAGCCACGCAGTTTTGATTA

CC

SecG_P2 TTGGGCTGGCGCGAGC

TAGGATATAAGCAAGCGC

SecG_seq TCTGATACAGCGTGACATGC

BdbB_P1 GGGTTCCTGGCGCGAGC

TGGCACAATGGCTAGTTTATTC

BdbB_P2 TTGGGCTGGCGCGAGC

ATTATATACATGTTGATTTTG

BdbB_seq TCTTGCCAACTCACTCCAC

BdbC_P1 GGGTTCCTGGCGCGAGCTATGCTTCCTGGGTTGT

G

BdbC_P2 TTGGGCTGGCGCGAGCTTCAGATTTTTCGCCTTTCA

GC

BdbC_seq TGCGGATTATGATGAAATC

SppA_P1 GGGTTCCTGGCGCGAGCCCATAAGGACATTATG

TCTC

SppA_P2 TTGGGCTGGCGCGAGCCTTCGCATAGAGATACA

TC

SppA_seq TCGGCGTCATTATGGAAAGC

YteJ_P1 GGGTTCCTGGCGCGAGC

Chapter 6. APPENDIX

182

TTCCTGCTTGATTGGCTC

YteJ_P2 TTGGGCTGGCGCGAGC

TTTGCGGTATAATTTTTCATGC

YteJ_seq AGAGTTAGAGCGCAATGACATC

SipS_P1 GGGTTCCTGGCGCGAGC

TGACTCTATGTATCCTACAC

SipS_P2 TTGGGCTGGCGCGAGC

ATTTGTTTTGCGCATTTCG

SipS_seq AGCAATTGTGATTGCTGTCG

SipT_P1 GGGTTCCTGGCGCGAGC

CACATTACATGACGGAGAAAGG

SipT_P2 TTGGGCTGGCGCGAGC

TTTTGTTTGACGCATTTCG

SipT_seq TCATCGCTGTTCTGCTGGC

PrsA_P1 GGGTTCCTGGCGCGAGC

AGGCGAGAAGTTTGAAGACC

PrsA_P2 TTGGGCTGGCGCGAGC

TTTAGAATTGCTTGAAGATG

PrsA_seq AAGGCAAAATCCGTGCAAGC

SpoIIIJ_P1 GGGTTCCTGGCGCGAGC

AGCTTCAACAGGAAACAATGGC

SpoIIIJ_P2 TTGGGCTGGCGCGAGC CTTTTTCTTTCCTCCGGC

SpoIIIJ_seq AGTTCGAAAGCGATGCAGGC

N-terminal SPA tag

fusion

SipS_1N cagCTCGAGATTGGAGGAAATCAGTTG

SipS_2N cagGCGGCCGCGCTGCTTGATCCTAATTTG

SipT_1N catCTCGAGCTAGGAGGAAACACTTTGACC

SipT_2N ataGCGGCCGCAAACGCCTGCTGTCCTAGG

Chapter 6. APPENDIX

183

SppA_1N gtgCTCGAGGGAGAAAAAATGAATGC

SppA_2N attGCGGCCGCTTGTTCCCTCCTACTTCGC

pSPANter_P1 GGAGATTCCTAGGATGGACTAC

pSPANter_P2 ACTGGTCTGATCGATGGGATGTC

Deletion mutants

Ko_ytej_P1 GGATTGCATTAGTGATTGCTCTGG

Ko_yteJ_P2 ACTATTGCCGATGATAAGCTGTATTGTTCCCTCC

TACTTCGCATAG

Ko_yteJ_P3 TGCGAAGTAGGAGGGAACAATACAGCTTATCAT

CGGCAATAGTTAC

KO_yteJ_P4 ACGGAAGTCCGGGTTATCGGGGCGATATAGGCG

CCAGCAAC

Ko_yteJ_P5 TTGCTGGCGCCTATATCGCCCCGATAACCCGGA

CTTCCGTC

Ko_yteJ_P6 TCTGATGCCTGTCGATCCGAC

ko_YteJ_P7 AGCTTCGGATTAGGCTCACTG

Ko_yteJ_P8 AGATACTCCAGCGCGACATG

KO_sppA_P1 TCATAAAAGGACAGGCAGCAGC

Ko_sppA_P2 AGGGTAACTATTGCCGATGATAAGCTGCCCAGA

GCAATCACTAATGC

Ko_sppA_P3 GGATTGCATTAGTGATTGCTCTGGGCAGCTTATC

ATCGGCAATAG

ko_sppA_P4 TGTCGCGTCCATTATTGTTCCCTCGGCGATATAG

GCGCCAGCAACC

Ko_sppA_P5 TTGCTGGCGCCTATATCGCCGAGGGAACAATAA

TGGACGCGAC

Ko_sppA_P6 GTTAACACACCTGTCACTGC

Ko_sppA_P7 TCTGCCATCCAGTATTTGTG

Chapter 6. APPENDIX

184

ko_sppA_P8 TCACGAGCCAATCAAGCAGG

ko_sppA.yteJ_P1 CGTGATTAACGGCCAGATTG

ko_sppA.yteJ_P2 CTATTGCCGATGATAAGCTGTTTTTCTCCTCCTT

TTTCCTAAACTC

ko_sppA.yteJ_P3 TAGGAAAAAGGAGGAGAAAAACAGCTTATCAT

CGGCAATAGTTACC

ko_sppA.yteJ_P4 GGACGGAAGTCCGGGTTATCGGGGCGATATAGG

CGCCAGCAAC

ko_sppA.yteJ_P5 TTGCTGGCGCCTATATCGCCCCGATAACCCGGA

CTTCCGTCCG

ko_sppA.yteJ_P6 AGCATTCTGATGCCTGTCGATC

ko_sppA.yteJ_P7 TTGGCCATAAGTGTACCACG

ko_sppA.yteJ_P8 CACTTCCTCTAGGAGCATTATCG

Transcriptional

sGFP fusions

Psig_P1 *(Rev) CAGGATCCGAATCGTTTCAGT

Psig_P2.2 *(Fw) ATGTCAAAAGGAGAAGAACT

Psig_P1_ext *(Rev)

TTTTTCTCCTCCTTTTTCCTAAACTCTCTCATTAT

ACGA

TTCAATATGAACAGGATCCGAATCGTTTCAGT

Psig_P2_ext *(Fw)

TTCATATTGAATCGTATAATGAGAGAGTTTAGG

AAAA

AGGAGGAGAAAAACATGTCAAAAGGAGAAGAA

C

Psig_P3 TCTGCCATCCAGTATTTGTG

Psig_P4 TTTTTCTCCTCCTTTTTCCT

Psig_P3_ext ACTGAAACGATTCGGATCCTGTCTGCCATCCAG

TATTTGTG

Psig_P4_ext GTTCTTCTCCTTTTGACATGTTTTTCTCCTCCTTT

TTCCT

Chapter 6. APPENDIX

185

PsigAW_P3_ext ACTGAAACGATTCGGATCCTGATCCCATTTCCCC

TTTGCTG

PsigAW_P4_ext AGTTCTTCTCCTTTTGACATTTTTTCTCCTCCTTT

TTCCT

PsigW_P3 ACTGAAACGATTCGGATCCTGGTTTGCCGGCAG

CCCGTTAAG

Translational sGFP

fusions

SG13_P1 TGGTCTGATCGCTCGGATACC

SG13_P2 ACATCACCATCCAATTCAACC

YteJ-GFP_P1 AACTAATAAGGAGGACAAACATGGACGCGACAT

ATGAAGA

YteJ-GFP_P2 TGAGCCGCCGCCGCCTGAGCCGCCGCCGCCTTT

GCGGTATAATTTTTCAT

SG13_YteJ-GFP_P3 GGCGGCGGCGGCTCAGGCGGCGGCGGCTCAAT

GTCAAAAGGAGAAGAACT

SG13_YteJ-GFP_P4 TCTTCATATGTCGCGTCCATGTTTGTCCTCCTTATT

AGTT

GFP-SppA_P1 GGCGGCGGCGGCTCAGGCGGCGGCGGCTCAAT

GAATGCAAAAAGATGGAT

GFP-SppA_P2 TGACTATTCTCAGATAAAGTCTACTTCGCATAG

AGATACATC

SG13_GFP-SppA_P3 GATGTATCTCTATGCGAAGTAGACTTTATCTGA

GAATAGTCA

SG13_GFP-SppA_P4 TGAGCCGCCGCCGCCTGAGCCGCCGCCGCCTTT

ATAAAGTTCGTCCATAC

GFPseq_P1 TGCTTCCAGATAACCACTACC

GFP seq_P2 ACCGTTAGTTGCATCACCTTC

sGFP_RBS_P1 AGCGATTAACTAATAAGGAG

sGFPSppA_P2.2 CAGGTCGACCTACTTCGCATAGAGATACATC

Chapter 6. APPENDIX

186

sGFP_P2.2 CGGTCGACTTATTTATAAAGTTCGTCCATAC

Screening and

sequencing

CsppA_P1 CCTAGGTCTGCCATCCAGTATTTGTG

CsppA_P2 CTCGAGTGTTCCCTCCTACTTCGCATAG

AmyE3_P1 CTCGTTACACCATCACTGTTCG

AmyE5_P2 TTCTCCAGTCTTCACATCGG

KOseq_P1 ACAGTTCTCAGTGGTGATCC

KOseq_P2 CTTGAACATTTGCTGGATGC

psweetCmFor CAGCTTATCATCGGCAATAG

psweetCmRev GGCGATATAGGCGCCAGCAACC

Nter-AmyE P1 AATTCTGCGTGACATCCCATCG

Nter-AmyE P2 TCGACATGGATGAGCGATGATG

Nter_SpecP1 TCATGAGCAATTTGATTAAC

Nter_SpecP2 TAGCCTAATTGAGAGAAGTTTC

Spec_seq TCTTTGCCAGAACTAATTGAGG

AmyE_up TGTATTCACTCTGCCAAGTTG

AmyE_down TAAGTCCCGTCTAGCCTTGC

His and SPA tag

fusion

Nter-HisP1 GCGAAATTAATACGACTCAC

Cter_HisP2 CTTGGCTCGACTTAATGGTG

SppAHis_P1 CTGCATATGAATGCAAAAAGATGGATTGC

SppA-CHis_P2 CTTCGCATAGAGATACATCATTC

SppANHis_P2 CTGCTCGAGCTACTTCGCATAGAGATACATC

YteJ_His_P1 TGGACGCGACATATGAAGAGTTAG

YteJ-CHis_P2 TTTGCGGTATAATTTTTCATGC

YteJ_NHis_P2 CTGCTCGAGTTATTTGCGGTATAATTTTTC

His_SYop-P1 TGAATGCAAAAAGATGGATTGCATTAG

Chapter 6. APPENDIX

187

SSppAHis_P1 ACGCATATGAACGGATCTCACATCACTGACG

His-SSppA_P1.35 CAGCATATGCAAACGGATCTCACATCACTGACG

His-SSppA_P1.53 CGCATATGGGCAGTCCCTCAAGTAAAATTGCC

FHis-SppA

TAGGAGGTAAAAAAAATGCACCATCACCAT

CACCATATGAATGCAAAAAGATGGATTGC

SPA_Rev CTGGTCGACCTACTTGTCATCGTCATCCTTG

YteJSPA-P2 GCCGCTAGCTTTGCGGTATAATTTTTCATGC

Overexpression

strains

pDR111-P1 TTGCTCGAGGGTAAATGTGAGC

Hspank_SppA_P2 CAGGTCGACCTACTTCGCATAGAGATAC

Hspank_SppA_P1 AGGAGGAGAAAAAATGAATGC

Hspank_YteJ_P1 CAGAAGCTTAGGAGGGAACAATAATGGACG

Hspank_YteJ_P2 CAGGTCGACTTATTTGCGGTATAATTTTTC

FsppA-K199A AGCGGGGCCCATGCGGACATTATGTCT

BsppA-K119A AGACATAATGTCCGCATGGGCCCCGCT

YteJ_LacA-P1 TGCGGATCCAGGAGGTAAAAAAAATGGACGCG

ACATATGAAGAG

YteJ_LacA-P2 CTGCCGCGGTTATTTGCGGTATAATTTTTC

XylR_seq TACTTGAGAGGAGACAGTAGAC

LacA_up TGCCGTCATCTTTATTATGCTGCA

LacA_down CCTGTTGAACTACATGCACTCCAC

RYteJ_93 CTGGTCGACTTACTGCCTGAAGTATTTTGTCATG

AG

RYteJ_129 CTGGTCGACTTAATCAATATACCGTCCAACCAC

RYteJ_152 CTGGTCGACTTACGCGATGTAATCATGAATCCC

RYteJ_55 CTGGTCGACTTAAAAGAGGGGAGAAACGAGCA

A

Chapter 6. APPENDIX

188

SppA-SeqP2 TCTGTGGTTATATCCGTCTGCACC

Table S 4. List of B. subtilis strains

Strain Genotype Source

GHBs 001 BSB1 (Trp+) strain Laboratory

stock

GHBs 006 BSB1 sipT::sipT-SPA lacImC;EryR This study

GHBs 008 BSB1 sipS::sipS-SPA lacImC;EryR This study

GHBs 010 BSB1 secG::secG-SPA lacImC;EryR This study

GHBs 012 BSB1 sppA::sppA-SPA lacImC;EryR This study

GHBs 014 BSB1 bdbC::bdbC-SPA lacImC;EryR This study

GHBs 017 BSB1 secDF::secDF-SPA lacImC;EryR This study

GHBs 018 BSB1 yteJ::yteJ-SPA lacImC;EryR This study

GHBs 020 BSB1 spoIIIJ::spoIIIJ-SPAlacImC;EryR This study

GHBs 023 BSB1 ∆yteJ::cm This study

GHBs 025 BSB1 ∆sppA::cm This study

GHBs 028 BSB1 ∆sppA-yteJ::cm This study

GHBs 029 BSB1 pCS74 This study



GHBs 032 BSB1 ∆yteJ::cm pCS74 This study



GHBs 035 BSB1 ∆sppA::cm pCS74 This study


Chapter 6. APPENDIX

189


GHBs 038 BSB1 ∆sppA-yteJ::cm pCS74 This study



GHBs 041 NCIB3610 (wt strain) Laboratory

stock

GHBs 043 NCIB3610 ∆yteJ::cm This study

GHBs 048 BSB1 amyE::Paw-GFP This study

GHBs 054 BSB1 amyE::Pw-GFP This study

GHBs 055 BSB1 amyE::SPA-sipS This study

GHBs 056 B.subtilis ATCC6633 (subtilin) Laboratory

stock

GHBs 059 BSB1 amyE::Phyper-spank yteJ spec This study

GHBs 061 BSB1 ∆yteJ::cm amyE::Phyper-spank yteJ spec This study

GHBs 062 BSB1 ∆sppA-yteJ::cm amyE::Phyper-spank yteJ spec This study

GHBs 064 BSB1 amyE::Phyper-spank yteJ-gfp spec This study

GHBs 065 BSB1 ∆sppA-yteJ::cm amyE::Phyper-spank yteJ-gfp spec This study

GHBs 066 BSB1 amyE::Phyper-spank sppA spec This study

GHBs 067 BSB1 ∆sppA::cm amyE::Phyper-spank sppA spec This study

GHBs 068 BSB1 ∆sppA-yteJ::cm amyE::Phyper-spank sppA spec This study

GHBs 069 BSB1 amyE::Phyper-spank gfp-sppA spec This study

GHBs 070 BSB1 ∆sppA-yteJ::cm amyE:: Phyper-spank gfp-sppA spec This study

GHBs 071 BSB1 amyE:: Phyper-spank sppA-yteJ spec This study

GHBs 072 BSB1 ∆sppA-yteJ::cm amyE:: Phyper-spank sppA-yteJ spec This study

GHBs 073 BSB1 amyE:: Phyper-spank sppA pCS74 spec neo This study


Chapter 6. APPENDIX

190


GHBs 076 BSB1 amyE:: Phyper-spank yteJ pCS74 spec neo This study



GHBs 079 BSB1 amyE:: Phyper-spank sppA-yteJ pCS74 spec neo This study



GHBs 085 BSB1 amyE:: Phyper-spank sppA-yteJ∆129-164 spec This study

GHBs 087 BSB1 amyE:: Phyper-spank sppA-yteJ∆152-164 spec This study

GHBs 089 BSB1 amyE:: Phyper-spank sppA-yteJ∆92-164SPA spec This study

GHBs 089 BSB1 amyE::Phyper-spank sppAK199A spec This study

Table S 5 List of E. coli strains

Strain Genotype Source

GHEc 002 E.coli pSPA_LIC+bdbB This study

GHEc 004 E.coli pSPA_LIC+secDF This study

GHEc 005 E.coli pSPA_LIC+secG This study

GHEc 007 E.coli pSPA_LIC+sdbC This study

GHEc 009 E.coli pSPA_LIC+sipS This study

GHEc 011 E.coli pSPA_LIC+sipT This study

GHEc 013 E.coli pSPA_LIC+sppA This study

GHEc 015 E.coli pSPA_LIC+spoIIIJ This study

GHEc 016 E.coli pSPA_LIC+yteJ This study

GHEc 026 E.coli MC1061 + pJ411_His-yteJ This study

GHEc 029 E.coli MC1061 + pJ411_His-sppA This study

Chapter 6. APPENDIX

191

GHEc 032 E.coli BL21 + pJ411_His-yteJ This study

GHEc 033 E.coli BL21 + pJ411_His-sppA This study

GHEc 043 E.coli MC1061 pDR111 yteJ This study

GHEc 046 E.coli MC1061 pDR111 yteJ-GFP This study

GHEc 047 E.coli MC1061 pDR111 sppA This study

GHEc 050 E.coli MC1061 pDR111 GFP-sppA This study

GHEc 052 E.coli MC1061 pDR111 sppA-yteJ This study

GHEc 054 E.coli 2566 pJ411 His-sppA This study

GHEc 055 E.coli 2566 pJ411 His-steJ This study

GHEc 056 E.coli 2566 pJ411 His-sppA-yteJ This study

GHEc 057 E.coli 2566 pSMG212 His-sppA-yteJ-TEV_FLAG This study

GHEc 058 E.coli 2566 pJ411 His-∆52SppA This study

Table S 6 List of plasmids

Plasmid Construction

Resistance

Source

E.coli B.subtilis

pGH001 pMutin SPA_Lic AmpR EryR Laboratory

stock

pGH002 pSPANter_Nter AmpR SpecR Laboratory

stock

pGH004 pMutin SPA_Lic with bdbB gene AmpR EryR This study

pGH006 pMutin SPA_Lic with secDF gene AmpR EryR This study

pGH007 pMutin SPA_Lic with secG gene AmpR EryR This study

pGH009 pMutin SPA_Lic with bdbC gene AmpR EryR This study

pGH011 pMutin SPA_Lic with sipS gene AmpR EryR This study

pGH013 pMutin SPA_Lic with sipT gene AmpR EryR This study

Chapter 6. APPENDIX

192

pGH015 pMutin SPA_Lic with sppA gene AmpR EryR This study

pGH017 pMutin SPA_Lic with spoIIIJ gene AmpR EryR This study

pGH018 pMutin SPA_Lic with yteJ gene AmpR EryR This study

pGH021 pMutin SPA_Lic with prsA gene AmpR EryR This study

pGH022 pCS74 NeoR From DSM



pGH025 pSG13 AmpR SpecR Laboratory

stock

pGH026 pSG13_PAW-GFP AmpR SpecR This study

pGH029 pSG13_PW-GFP AmpR SpecR This study

pJ411_6His KanR Laboratory

stock

pGH032 pJ114_His-YteJ KanR This study

pGH035 pJ114_His-SppA KanR This study

pGH037 pSPANter_Nter SPA-Sips AmpR SpecR This study

pGH032 pJ114_His-SSppA KanR This study

pJ114_His-operon KanR This study

pGH042 pDR111 AmpR SpecR Laboratory

stock

pGH043 pDR111 yteJ AmpR SpecR This study

pGH046 pDR111 yteJ-GFP AmpR SpecR This study

pGH047 pDR111 SppA AmpR SpecR This study

pGH050 pDR111 GFP-SppA AmpR SpecR This study

pGH052 pDR111 operon AmpR SpecR This study

pGH053 pJ114 His-SppA-YteJ KanR This study

Chapter 6. APPENDIX

193

pSMG212 KanR Laboratory

stock

pGH054 pSMG212 His-SppA-YteJ-

TEV_FLAG

KanR This study

Figure S 1 YteJ predicted structure by Protter. Extra- extracellular compartment; Intra- cytosolic

compartment; The structure shows iYteJ amino acid composition, the amino acids are represented by the unique

letter symbol (Omasits et al. 2014).

Chapter 6. APPENDIX

194

Figure S 2. Purification of SppA, YteJ, SppA/YteJ and SppAK199A by Gel Filtration. (A)

Chromatograph obtained with the gel filtration of SppA, YteJ, SppA/YteJ and SppAK199A samples ;

(B) The protein samples were injected in a Superdex 200 10/300 GL gel filtration and all the fractions

from 4-23 were analysed by SDS-PAGE. The two bands 40kDa and 25KDa are highlighted in the

protein ladder.

Chapter 6. APPENDIX

195

Figure S 3 Pulse chase assays for sppA, yteJ and sppA yteJ deletion mutants and complemented strains. All the strains were transformed with pHTK10 (AmyQ secretion),as such were grown at MBU supplemented

with Kanamycin. (A) pulse chase gel results. P-pre-protein. M-mature protein; (B,C) Graphic representation of

the AmyQ processing results.

Chapter 6. APPENDIX

196

Figure S 4. The growth of B. subtilis sppA and yteJ deletion mutant strains in the presence of different

concentrations of LP9. The BSB1, ∆sppA and ∆yteJ deletion strains grown in LB in a 96-well plate, where the

OD600 was measured on specific time points. The different graphs represent the growth of the strains in the

presence of different concentrations (0 µg, 50 µg, 75 µg, 100 µg, 150 µg and 200µg) LP9, labelled on the Top of

the graph. Colours, on the picture, indicate the wild type or mutated strains; Black arrow represents the time

point of induction with LP9. Data from our collaborators at the Prof. Jan Maarten van Dijl.

Chapter 6. APPENDIX

197

Figure S 5. The growth of B. subtilis sppA and yteJ overexpressing mutant strains in the presence of

different concentrations of LP9. The BSB1, ∆sppA and ∆yteJ deletion strains grown in LB in a 96-well plate,

where the OD600 was measured on specific time points. The different graphs represent the growth of the strains

in the presence of different concentrations (0 µg, 50 µg, 75 µg, 100 µg, 150 µg and 200µg) LP9, labelled on the

Top of the graph. Colours, on the picture, indicate the wild type or mutated strains; Black arrow represents the

time point of induction with LP9. Data from our collaborators at the Prof. Jan Maarten van Dijl.

Chapter 6. APPENDIX

198

6.2 UCB Secondment: Purification and characterization

of a maltogenic α-amylase from Bacillus

stearothermophilus

A maltogenic amylase from B. stearothermophilus was overexpressed in B. subtilis at the

DSM company. Dr. Van Rij, one of our collaborators from the Protein Factory consortium, sent us a

crude medium supernatant sample from B. subtilis AmyM-secreting cells to UCB Company, at

Slough. For my secondment in this company, I started a collaboration with the group of Dr. David

Humphreys at UCB, which project objective it is the purification of the AmyM protein. This work has

been developed under the supervision of Dr. Emma Dave and Dr. Neesha Dedi.

This AmyM protein has a molecular weight of 75.2KDa, but in SDS-Page gels run below the

expected, around 66KDa as is possible to see on the Figure S 6. Its theoretical isoelectric point is 5.38.

This calculation is based on the protein sequence. According, to this information and to the

purification process published by Cha et al. in 1998, we begun a small scale purification with a

volume of 1ml of the supernatant sample (20mg/ml). The sample was diluted up to 2ml, applied to a

Q-Sepharose resin (a strong anion exchanger) equilibrated in 20mM Tris HCl pH7,5 and 5mM

CaCl2 and eluted with a linear gradient of NaCl from 100mM up to 1M. Consequently, as is

possible to see on the Figure 1 below, the protein was collected from the flow-through. The protein

did not bind to the resin beads, which may be due to a high salt concentration of the supernatant

buffer, making impossible the exchange of ions.

Chapter 6. APPENDIX

199

Figure S 6. AmyM supernatant purification. The AmyM sample was purified by anionic exchange

(Q-Sepharose resin). The different eluted samples and fractions were analysed by 10% SDS-PAGE gel. The

description for each lane shown at the right side of the picture.

This experiment was repeated but the result obtained was the same. The expiry date can may

be the reason why the AmyM sample is not binding to the Q-rresin. To test this hypothesis, the

concentrated enzyme sample was applied to a new Q-resin charged with two columns volumes of 20

mM Tris HCl and 1 M NaCl, washed with and equilibrated with 20 mM Tris HCl pH 7.5 and 5 mM

CaCl2. Like the previous experiments the protein was collected from the run-trough. This fraction was

reapplied to a Size exclusion Hitrap 16/60 Superdex 200 column, that allows the separation of the

protein and contaminants based on their size. The protein was eluted as a single peak with an

apparent molecular weight of 75 KDa, according with the molecular weight predicted. Although, the

protein has been eluted in a single peak the analysis by SDS-PAGE gel of each fractions

correspondent to the protein elution showed that some contaminants were eluted at the same peak

(Figure S 7).

The observation of some contaminants in the protein sample after the gel filtration raised the

question, is the pH buffers used for the anionic exchange chromatography correct?

Therefore, to test this hypothesis the protein sample was subjected to a new anion exchange

chromatography at 4 °C in a 1 ml Hitrap HP-Q column equilibrated in 20 mM Tris HCl pH 8.5 and 5

mM CaCl2. The protein was eluted with a linear gradient of NaCl from 0 mM up to 1 M. Through

the analysis of the UV value measured at 280 nm it was possible to see that the protein did not bind

to the resin and was eluted during the flow-through. The flow-through sample was concentrated

(with an Amicon Ultra-15 column) and analysed by SDS-PAGE gel, were we could confirm the

presence of AmyM protein at the flow-through.

Chapter 6. APPENDIX

200

Figure S 7. AmyM Size exclusion purification results. The protein sample was size-fractionated at 4 °C using

a Hitrap 16/60 Superdex 200 column equilibrated with 20 mM Tris HCl pH 7.5, 5 mM CaCl2 and 100 mM NaCl

buffer and eluted at a flow rate of 2 ml/min. The legend of the loaded samples is described at the right side of the

figure.

Consequently, it was performed a cationic exchange chromatography at 4 °C in a 1 ml Hitrap

HP-S column equilibrated in 20 mM Tris HCl pH 7.5 and 5 mM CaCl2. The protein was eluted with a

linear gradient of NaCl from 0 mM up to 1 M. The measurements of the UV values at 280 nm

showed that the protein had not bound to the resin and, once more, the protein was eluted with the

flow-through.

Then the AmyM sample was reapplied to a Hitrap 26/60 Superdex 200 column equilibrated

with TBS and 5 mM CaCl2 and eluted at a flow rate of 2 ml/min for 5 hours. This column allows a

better separation of the different fractions of the sample since have a bigger size. The protein was

eluted as a single peak with an apparent molecular weight of 75 KDa. However, the sample fractions

correspondent to the protein elution showed the contaminants were kept.

Although the theoretically pI be 5.38 (Dauter et al. 1999; Cha et al. 1998; Park et al. 2013)

it is possible that this is not the real value, what could explain the why the protein is not binding to

the anionic or cationic resin of the exchange chromatography process. In 1984, H. Outtrup and B. E.

Norman studied the B. stearothermophilus maltogenic amylase and determined by thin layer gel-

electrofocusing that this protein has a pI of 8.5. To confirm the correct pI value we measured the pI

of our sample by iCE3 Capillary IEF System. This is a HPLC-like method that performs free solution

Isoelectric Focusing (IEF) in a capillary column, in which a pH gradient is created to separate and

focus the proteins according to their pI. The all process is monitored in real time by a whole-column

UV detector. Before run, the samples are premixed with carrier ampholytes, additives and pI markers.

The pI measured for our sample was 8.35. To complement the pI measurement a molecular weight

analysis was performed by time-of-flight mass spectrometry (LC- QTOF). The used technique allows

the obtainment of the protein molecular weight thought the determination of an ion mass-to-charge

ratio that is measured according to the time the ion takes to reach a detector at a known distance

(Sigaud et al. 2016; Kernphysik, Darmstadt, and Wien 1999; Beauchemin 2009). The measure

Chapter 6. APPENDIX

201

molecular weight was 75157 Da. This value is in good agreement with the information received from

DSM with the details of the supernatant sample but is slightly different from those reported previously

(Cha et al. 1998; Outtrup 1984).

The real pI value (8.35) of the AmyM sample can explain the problems that we had with the

binding of the protein to the cationic and anionic resins, because the range of pH tested was from 7.5

up to 8.5 and to be possible the exchange of ions the pH value used should be two values above or

below of the protein pI value (Svensson 1961) .

Furthermore, to avoid high pH values in which the protein is denatured we tested a cationic

exchange chromatography at pH 6.5. The buffer of a 2 ml supernatant sample was exchange to 50 mM

MES pH 6.5 and 50 mM NaCl with the PD-10 Desalting columns GE Health Care. The sample was

then diluted up 20 ml and applied to a cationic a 1 ml Hitrap HP-S column equilibrated in 50 mM

MES pH 6.5 and 50 mM NaCl. The protein was eluted with a linear gradient of NaCl from 50 mM up

to 1 M. During the collection of the flow-through fraction it was possible to observe an increase of

the UV values at 280 nm what normally indicates that the protein is not binding to the column resin,

but because it is not a high value as the values previous measured it can be some contaminants that

did not bound to the resin. Therefore, a 50 mM MES pH 6.5 buffer with 2 M NaCl was loaded into

the purification system to create a gradient of NaCl up to 2 M. This additional step has the objective of

elute the sample in the case that it binds tightly and it is necessary to have higher concentrations of salt

to elute the protein. However, it was not the case and the protein was collected in the flow-through

fraction once again.Since the previous purifications were not successful we procedure to a new anionic

exchange chromatography with an increase of the pH buffer up to 9,5 (Dauter et al. 1999). A 2 ml

protein sample from the previous experiment Flow-through fractions sample was buffer exchanged to

20 mM Tris-HCl pH 9.5 with the PD-10 Desalting columns. The sample applied to an anionic 1 ml

Hitrap HP-Q column equilibrated in 20 mM Tris-HCl pH 9.5. The protein was eluted with a linear

gradient of NaCl from 50 mM up to 2 M. Following the UV measured values during the

chromatography process it is possible to see that the protein bound to the resin and it was eluted as a

single peack. However, after SDS-PAGE analysis the protein sample had yet other bands besides the

band corresponding to the AmyM molecular weight as shown at Figure S 7. These bands could

represent contaminants or cleaved protein, because by either processes, by ions exchange process or

by molecular exclusion size this, it seems impossible to rid of the contaminant bands. By the

comparison of the abundance of the AmyM and the other specimens presents in the sample is possible

to know that the AmyM sample is clean enough for the rabbit Immunization. This purification step

was applied to all the remaining protein sample. After, all the sample passed through a endotoxin free

removal protocol and were send to the rabbit Immunization department.

Université Paris-Saclay

Espace Technologique / Immeuble Discovery

Route de l’Orme aux Merisiers RD 128 / 91190 Saint-Aubin, France

Titre : Caractérisation du complexe membranaire impliquant la signal peptide peptidase

SppA et YteJ chez Bacillus subtilis.

Mots clés : Sécretion protéique, complexe protéique membranaire, protéase, résistance aux

lantibiotiques, contrôle qualité

Résumé : Chez Bacillus subtilis nous avons identifié un complexe protéique membranaire impliquant

une protéine inconnue, YteJ, et une autre protéine membranaire, SppA, une signal peptide peptidase

également impliquée dans la résistance aux peptides antibactériens de la famille des lantibiotiques.

Après délétion des gènes correspondant, nous avons montré que les deux protéines sont impliquées dans

cette résistance. Dans la souche ∆sppA, la surexpression ectopique de SppA a non seulement restauré la

résistance, mais elle a également induit la formation de cellules allongées, un phénotype supprimé par

la surexpression simultanée de YteJ. L'expression de versions tronquées de YteJ a mis en évidence le

rôle inhibiteur d'un domaine spécifique de YteJ. Enfin, des études biochimiques in vitro ont confirmé

que l'activité de la protéase SppA était fortement réduite par la présence de YteJ, confirmant l'hypothèse

d'une inhibition par YteJ. Nos études in vivo et in vitro ont montré que YteJ, via l'un de ses domaines,

agit comme régulateur négatif de l'activité protéase de SppA dans ce complexe. En conclusion, nous

avons montré que le complexe SppA/YteJ est impliqué dans la résistance aux lantibiotiques à travers

l’activité protéase de SppA, elle-même régulée par YteJ.

Title : Towards the understanding of the function and regulation of a membrane complex

involving SppA and YteJ in Bacillus subtilis.

Keywords : Bacillus subtilis, Protein secretion, Membrane protein complex, Protease, Lantibiotic

resistance, Quality control

Abstract: We have identified a membrane protein complex of Bacillus subtilis involving an

unknown protein, YteJ, and SppA, a membrane protein first described as a signal peptide

peptidase and later shown to be also involved in the resistance to antibacterial peptides of the

lantibiotic family. Using deletion mutant strains, we showed that both proteins are involved in

this resistance. In the ∆sppA strain, the ectopic overexpression of SppA not only restored the

resistance, it also induced the formation of elongated cells, a phenotype suppressed by the

simultaneous overexpression of YteJ. Furthermore, the expression of truncated versions of YteJ

pinpointed the inhibitory role of a specific domain of YteJ. Finally, in vitro biochemical studies

showed that SppA protease activity was strongly reduced by the presence of YteJ, supporting

the hypothesis of an inhibition by YteJ. Our in vivo and in vitro studies showed that YteJ, via

one of its domain, acts as a negative regulator of the protease activity of SppA in this complex.

In conclusion, we have shown that SppA/YteJ complex is involved in lantibiotic resistance

through the protease activity of SppA, which is regulated by YteJ.

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